How to become a PCR pro

HOW TO BECOME A PCR PRO

The polymerase chain reaction (PCR) is a key life sciences technique. It has been used in

molecular biology – including molecular diagnostics – for many years, and a number of different

types, for example, RT-PCR, qPCR, vPCR and ddPCR have been developed over time.

Today, PCR is a vital tool for the detection of pathogens, such as the SARS-CoV-2 virus, and

is essential for genotyping and NGS library preparation. However, PCR is well known for being

difficult to run successfully and several parameters must be considered when planning the PCR

protocol.

We have therefore compiled this eBook – consisting of in-depth educational articles, relevant

app notes and customer testimonials – to help you understand how PCR works, and what needs

to be considered to perform effective PCR reactions. We also demonstrate how our solutions

can help you to enhance the throughput of your lab, and become a PCR pro in no time.

Dr Éva Mészáros

Application Specialist

eva.meszaros@integra-biosciences.com

Anina Werner

Content Manager

anina.werner@integra-biosciences.com

FOREWORD

TABLE OF CONTENTS

CHAPTER 1: What you need to know about PCR

1.1 The complete guide to PCR 2

1.2 Simple PCR tips that can make or break your success 15

1.3 Setting up a PCR lab from scratch 20

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work 24

1.5 How to design primers for PCR 32

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 38

CHAPTER 3: Application notes

3.1 Efficient and automated 384 well qPCR set-up with the ASSIST PLUS pipetting robot 42

3.2 Automated RT-PCR set-up for COVID-19 testing 49

3.3 Increase your sample screening and genotyping assay throughput with the VOYAGER 57

adjustable tip spacing pipette

3.4 PCR product purification with QIAquick® 96 PCR Purification Kit and the 61

VIAFLO 96 handheld electronic pipette

3.5 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 66

VIAFLO 96

3.6 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 72

ASSIST PLUS

CHAPTER 4: Customer testimonials

4.1 INTEGRA pipettes – the obvious choice for start-up PCR labs 81

4.2 A better qPCR pipetting experience 83

4.3 COVID-19 – Accelerate your PCR set-up 85

4.4 Reducing protocol time for PCR using 96 channel pipette 86

CHAPTER 5: Conclusion 88

CHAPTER 6: References 89

2

CHAPTER 1:

What you need to know about PCR

In this chapter, we will cover topics such as PCR’s fascinating history, its mechanism and

different variations, and techniques for troubleshooting common issues you may encounter.

We’ll also go through tips for establishing a PCR lab, as well as a comprehensive overview of all

things related to qPCR and primer design.

1.1 The complete guide to PCR

Polymerase chain reaction (PCR) methods have been carried out in labs around the world since

the 1980s, opening the door for an array of new applications, such as genetic engineering,

genotyping and sequencing. In this article, we take a deep dive into this fascinating technique

by explaining its mechanism, exploring its history, looking into the different types of PCR,

discussing troubleshooting tips and much more.

CHAPTER 1: What you need to know about PCR

3

What is PCR?

The polymerase chain reaction (PCR) is a fast and inexpensive technique for amplifying a DNA

sequence of interest. It consists of three steps:

• Denaturation: The sample is heated to separate the DNA into two single strands.

• Annealing: The temperature is lowered to allow primers to anneal to specific single-stranded

DNA segments, flanking the sequence to be amplified.

• Extension: The temperature is raised to the optimum working temperature of the polymerase

enzyme, which then makes a complementary copy of the DNA sequence of interest.

One such repetition or 'thermal cycle' theoretically doubles the amount of the DNA sequence of

interest present in the reaction. Typically, 25 to 40 cycles are performed – resulting in millions

or even billions of DNA copies – depending largely on the amount of DNA in the starting sample

and the number of amplicon copies needed for post-PCR applications.

The three steps of a PCR reaction are carried out automatically by a thermal cycler, but can

only be successful if the master mix has been correctly prepared. The following sections

explain the components that make up the master mix and how they interact with the template

DNA during thermal cycling.

PCR master mix components

The PCR master mix consists of six components:

• PCR-grade water: Certified to be free of contaminants, nucleases and inhibitors.

• dNTPs: Containing equal concentrations of the four nucleotides (dATP, dCTP, dGTP and

dTTP), which are the 'building blocks' to create complementary copies of the DNA sequence

of interest.

• Forward and reverse primers: Short, single-stranded DNA sequences that anneal

specifically to the plus and minus strands of the template DNA, flanking the sequence to

be amplified. For some assays – such as protocols amplifying much-studied genes or DNA

sequences of common bacteria – ready-to-use primers can be purchased. However, many

experiments require custom PCR primers tailored to the region of interest of the template DNA

and the reaction conditions.

CHAPTER 1: What you need to know about PCR

4

• DNA polymerase: Taq-polymerase is the most commonly used enzyme for PCR reactions.

It uses dNTPs to create complementary copies of the DNA sequence of interest. For some

applications, such as mutagenesis, Taq-polymerase is not accurate enough and the use

of high fidelity polymerases is recommended. Just like Taq-polymerase, they sometimes

add an incorrect nucleotide when replicating the template DNA but, as they have a 3' to

5' exonuclease activity, they 'proofread' the newly synthesized strands and correct any

mistakes.1 This proofreading step is highly beneficial for accuracy but it also slows down PCR

reactions, and high fidelity polymerases (also called slow polymerases) therefore need about

twice the time of Taq-polymerase to create a complementary DNA strand. The most popular

high fidelity DNA polymerase is Pfu-polymerase.2

• Buffer: Provides a suitable environment for the DNA polymerase, with a pH between 8.0

and 9.5.3

• Magnesium chloride: Increases the activity of the DNA polymerase and helps primers

to anneal to the template DNA for a higher amplification rate.4 This cofactor is sometimes

included in the buffer in a sufficient concentration.5

The template DNA , which may be genomic DNA (gDNA), complementary DNA (cDNA) or

plasmid DNA (pDNA), is then added after master mix preparation.

The 3 steps of PCR

After preparing the PCR master mix and adding the template DNA samples to it, you can load

your reaction tubes, PCR strips or microplates into the thermal cycler. They will then go through

the following steps:

• Denaturation: The thermal cycler first heats the reaction mix to 95-98 °C to denature the

template DNA, separating it into two single strands. Depending on your sample, this usually

takes 2-5 minutes during the first thermal cycle, and 10-60 seconds for subsequent cycles.

• Annealing: When the temperature is lowered, the primers anneal to the sequences flanking

the template DNA region of interest. Depending on the sequence and melting temperature of

your primers, this step usually takes 30-60 seconds, and the optimal annealing temperature

typically lies between 45 and 60 °C.

• Extension: The temperature is increased to 72 °C, which is the ideal working temperature

for the Taq-polymerase. Depending on the synthesis rate of your polymerase, and the length

of the target sequence, it usually takes 20-60 seconds to create complementary copies

of the DNA sequence of interest.6 After approximately 25-40 cycles – depending on the

amount of DNA present at the start, and the number of amplicon copies needed for post-PCR

applications7 – the last extension step should be extended to 5 minutes or longer, allowing the

Taq-polymerase to finish the synthesis of uncompleted amplicons.5 If you can't immediately

take your samples out of the thermal cycler after the final extension step because you're busy

with other experiments, program it to cool your samples to 4 °C. For overnight runs where you

CHAPTER 1: What you need to know about PCR

5

leave your samples in the thermal cycler for hours after the final extension step, you should

opt for a holding temperature of 10 °C instead of 4 °C, as it causes less wear and tear on your

machine.

As shown in the image above, the amount of PCR product theoretically doubles at every

thermal cycle, leading to an exponential increase of PCR product. However, in reality, the phase

of exponential amplification eventually levels off and reaches a plateau because the reagents

have been consumed and the DNA polymerase activity decreases.

The different types of PCR

After performing a standard PCR reaction, you can determine the concentration, yield and

purity of the amplified DNA sequences using gel electrophoresis, spectrophotometry or

fluorometry. However, you can’t determine the amount of template DNA present in a sample

before amplification using standard PCR. If this is a requirement for your experiment, you have

to perform a qPCR reaction.

qPCR

qPCR – also called real-time PCR, quantitative PCR or quantitative real-time PCR – is a

technique used to detect and measure the amplification of target DNA as it is produced.

In contrast to conventional PCR reactions, qPCR requires a fluorescent intercalating dye

or fluorescently-labeled probes, and a thermal cycler that can measure fluorescence and

calculate the cycle threshold (Ct) value. Typically, the fluorescence intensity increases

proportionately to the concentration of the PCR product being formed, measuring quantities

of the target in real time.

CHAPTER 1: What you need to know about PCR

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qPCR can be divided into dye-based methods (e.g. SYBR® Green) and probe-based methods

(e.g. TaqMan®).

RT-PCR and RT-qPCR

Another limitation of standard PCR is that it can only be used to amplify DNA sequences. If you

want to amplify RNA target sequences, you have to use RT-PCR.

RT-PCR

vPCR

Reverse transcription PCR (RT-PCR) is used to amplify RNA target sequences, such as

messenger RNA or RNA virus genomes. This type of PCR involves an initial incubation of

the RNA samples with a reverse transcriptase enzyme and a DNA primer – comprising

sequence-specific oligo (dT)s or random hexamers – prior to the PCR amplification.

For viability PCR (vPCR), each sample needs to be split into two aliquots. One aliquot is

incubated with a photoreactive intercalating dye that is unable to diffuse through intact cell

membranes of live cells. This means that it only intercalates into the DNA of dead cells. When

this aliquot is subsequently treated with a blue light, the dye binds irreversibly to the DNA. Both

aliquots are then subject to DNA purification and qPCR amplification. If they exhibit similar

qPCR signals, the target microorganisms in the sample are mostly viable. If the dye-treated

aliquot exhibits a weaker signal, the target microorganisms are mostly dead. vPCR is an

important technique in diagnostics, agriculture and food safety.

You can also perform a qPCR reaction instead of executing a standard PCR reaction after

the reverse transcription step, which produces cDNA from RNA. This PCR variant is called

RT-qPCR.

vPCR

The third limitation of standard PCR is that it cannot distinguish between the DNA of viable

and non-viable cells. You should use vPCR if this is important to your application, for example,

because you want to know if the infectious microorganisms in a clinical sample are dead or

alive.

CHAPTER 1: What you need to know about PCR

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ddPCR

Digital droplet PCR (ddPCR) is another relatively new type of PCR. It uses fluorescently labeled

probes to detect DNA sequences of interest, and a water-oil emulsion system to split each

sample into about 20,000 nanoliter-sized droplets. After amplification, every droplet of the

sample is analyzed on its own. Droplets that contain at least one DNA sequence of interest emit

a fluorescent signal – and are consequently positive – while droplets without the DNA sequence

of interest don't fluoresce, and are therefore negative. Using the Poisson distribution, you can

then determine the concentration of the DNA sequence of interest in the original sample by

analyzing the ratio of positive to negative droplets for absolute quantification.8

An advantage of ddPCR compared to qPCR is that it's more precise. While qPCR can detect

two-fold differences in DNA target sequence variation, e.g. discriminate 1 copy from 2 copies

of a gene, ddPCR can discriminate 7 copies from 8 copies, which means that it can detect

differences as small as 1.2-fold.9 On top of that, ddPCR is better suited for multiplexing assays

if you want to determine the ratio of low abundance to high abundance DNA sequences of

interest, such as rare mutations on wild type backgrounds. When using qPCR, the fluorescent

signal from the high abundance sequences can dominate and obscure the signal from the

low abundance sequences. This risk is ruled out with ddPCR, as each droplet behaves as

an individual PCR reaction and contains either zero, one or, at most, a few sequences of

interest.10,11

ddPCR

CHAPTER 1: What you need to know about PCR

8

Due to these advantages, ddPCR is often preferred over qPCR for the detection of mutations

and SNPs (single nucleotide polymorphisms), allelic discrimination, gene expression studies,

and the analysis of copy number variations.12

Hot start PCR

If your PCR reaction results in non-specific amplification, you can try to increase the reaction

specificity using a hot start polymerase. This enzyme remains inactive during master mix

preparation and sample addition at room temperature, eliminating the risk that unintended

products and primer dimers are formed during PCR set-up.13

Nested and semi-nested PCR

Nested or semi-nested PCR are alternatives to hot start PCR that increase reaction specificity.

Nested PCR uses two sets of primers and two successive PCR reactions. The first set of

primers is designed to amplify a DNA sequence slightly longer than the sequence of interest.

During the second PCR reaction, the second set of primers that is specific to the sequence of

interest anneals to the amplicons of the first PCR reaction and helps to amplify the sequence of

interest.14,15

Nested PCR

CHAPTER 1: What you need to know about PCR

9

Semi Nested PCR

Semi-nested PCR works similarly to nested PCR. During the first PCR reaction, one primer

anneals to the sequence of interest and the second primer to a region flanking the sequence of

interest. This primer is then replaced with a second primer annealing to the region of interest

during the second PCR reaction.

The idea behind nested and semi-nested PCR is that, if non-specific products were amplified

during the first PCR reaction, these will not be amplified during the second PCR reaction, as the

primers cannot anneal to them.

Touchdown PCR

A third type of PCR developed to increase reaction specificity is touchdown PCR. The assay

set-up for touchdown PCR is identical to the set-up for standard PCR. The only difference lies in

the annealing step. During the first thermal cycle, the annealing temperature should be several

degrees above the optimal primer annealing temperature, then be lowered by 1-2 °C every

second cycle.16 These high temperatures during the first cycles avoid PCR primers forming

primer-dimers or binding to regions outside the DNA sequence of interest. The downside is

that the PCR primers don't all sufficiently bind to the template DNA, which leads to low yields.17

However, this can be tolerated, as the low yield of specific amplicons is then exponentially

amplified with every thermal cycle that is performed at the optimal annealing temperature.

CHAPTER 1: What you need to know about PCR

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The history of PCR

As we've shown, there are many different types of PCR, and some of them have only recently

been developed. However, the foundation for PCR was laid in the 1950s:

• In 1953, James Watson and Francis Crick discovered the double-helix structure of DNA, and

suggested that there might be a possible copying mechanism for DNA.

• Four years later, Arthur Kornberg identified the first DNA polymerase that was able to copy the

template DNA, although only in one direction.

• In 1971, Gobind Khorana and his team started to work on DNA repair synthesis. Their

technique used DNA polymerase repeatedly, but employed only a single primer template

complex, which did not allow exponential amplification.

• At the same time, Kjell Kleppe from Khorana's lab proposed a two primer system that would

double the amount of DNA in a sample, but no one actually conducted the experiment to

find out whether it worked. The reason for this was probably that there was not yet a DNA

polymerase that could withstand the high temperatures of the denaturation step. This means

that they would have had to add a fresh dose of enzyme after every thermal cycle.

• In 1983, Kary Mullis, working at Cetus Corporation, added a second primer to the opposite

strand, and realized that repeated use of DNA polymerase triggers a chain reaction that will

amplify a specific DNA sequence, thus inventing PCR. The patent got approved in 1987, and

he won the Nobel Prize in Chemistry six years later.

• In 1976, the thermostable enzyme Taq-polymerase – which is typically used in PCR today

– was first isolated from the bacterium Thermus aquaticus, which had been discovered in a

hot spring of Yellowstone National Park in 1969. When it was finally incorporated into PCR

workflows in 1988, it removed the need to add a new dose of enzyme after every thermal

cycle, paving the way for the invention of automated thermal cyclers.18,19,20

CHAPTER 1: What you need to know about PCR

11

PCR troubleshooting

One of the most important troubleshooting mechanisms is to always include positive and

negative control samples.

If the sequence of interest wasn't amplified in your positive control sample, your master mix,

template DNA or thermal cycler could be the source of the problem:

• Master mix: Have you added the right volume and concentration of each reagent, and have

you cooled your reagents during master mix preparation?

• Template DNA: Have you run an agarose gel to ensure that your template DNA isn't

degraded? Is your template DNA pure enough and, if not, have you purified it?

• Thermal cycler: Is the number of thermal cycles sufficient for your assay? Have you

programmed the device correctly, and is it calibrated to ensure that it performs the reaction

steps at the right temperatures?

If the sequence of interest was amplified in your negative control sample, one or more

components of your master mix is contaminated. PCR reactions are very sensitive, and create

large number of copies of DNA sequences from minute amounts of starting material, so

contamination is a common issue. To prevent it, the right lab set-up is crucial.

CHAPTER 1: What you need to know about PCR

12

Lab set-up

Ideally, your PCR lab should have two rooms, each divided into two areas. The first room should

be exclusively used for pre-PCR activities, and divided into a master mix preparation area and a

sample preparation area. The second room should have a dedicated area for amplification, and

another one for product analysis.

If you’re lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible. In addition to the spatial separation, you could also consider setting up your PCR

reactions in the morning, and performing the amplification and analysis steps in the afternoon.

Temporally separating the different steps of your PCR reactions may limit your flexibility and

make you lose some time, but lowers the risk of aerosols with high DNA concentrations from the

analysis area contaminating your master mix and samples in the pre-PCR area.

On top of these precautionary measures, you should always work in biosafety cabinets or

laminar flow hoods when setting up your PCR reactions, use different sets of pipettes for master

mix preparation, sample preparation and analysis, and make sure that you use filter tips and

consumables that are free of DNase, RNase and PCR inhibitors.

Specificity

Another major PCR challenge is specificity. As explained before, it can be improved using hot

start, nested, semi-nested or touchdown PCR. A further option to prevent the amplification of

regions outside the DNA sequence of interest, as well as the formation of secondary structures,

is to redesign your primers.

CHAPTER 1: What you need to know about PCR

13

Use this checklist to see whether your primers meet all the requirements:

• Are your primers between 18 and 24 bp long?

• Is your target sequence length between 100 and 3000 bp for standard PCR assays, or 75

and 150 bp for qPCR assays?

• Do your primers have melting temperatures between 50 and 60 °C, and within 5 °C of

each other?

• Have you performed a gradient PCR to determine the optimal annealing temperature?

• Does the GC content of your primers lie between 40 and 60 %?

• Have you avoided runs or repeats of four or more bases or dinucleotides?

• Have you made sure that your primers are not homologous to a template DNA sequence

outside the region of interest?

• Have you checked that your primers can't form stable secondary structures?

PCR equipment

The most important PCR instrument is certainly the thermal cycler but, as the right pipetting

devices can help create faster and more efficient workflows with fewer errors, we'll also look at

a few different liquid handling options in this section.

Thermal cyclers

Before the development of thermal cyclers, scientists had to manually move their samples

between water baths of different temperatures. The first thermal cycler prototype called 'Mr.

Cycle' also used water baths to heat and cool the samples, and was developed by engineers

at Cetus Corporation, where Kary Mullis worked when he invented PCR.21 Today's instruments

work with electric heating and refrigeration units, and many different models with various

additional features are available.

For standard PCR, a thermal cycler that can heat and cool your samples to the required

temperatures might be sufficient to complete the different reaction steps. However, your

thermal cycler will need additional properties – such as gradient capability or an integrated

fluorometer – if you want to perform gradient PCR assays to optimize primer annealing

temperatures, or qPCR assays to determine the amount of template DNA present in a sample

before amplification.

CHAPTER 1: What you need to know about PCR

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Pipettes

While the thermal cycler is the star of PCR labs, the right pipettes help you to process more

samples in less time, while ensuring maximal accuracy and precision. Electronic pipettes

offering a Repeat Dispense mode, for example, are a great option to boost the efficiency of

aliquoting master mix into an entire well plate. Adjustable tip spacing pipettes, paired with low

dead volume reagent reservoirs, can be a useful alternative to single channel pipettes when

transferring reagents and samples between different labware formats. And, if you want to

significantly cut your PCR set-up and purification time, pipetting robots or 96 and 384 channel

pipettes might be the right tool for you.

Conclusion

We hope that this article has been useful in helping you understand the mechanisms behind

the different types of PCR, and has shown you different ways to avoid contamination and nonspecific

amplification.

CHAPTER 1: What you need to know about PCR

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1.2

Since the outbreak of the COVID-19 pandemic, PCR is on everybody's lips. However, only

people working in the lab know how difficult it can be to get the desired results using this wellestablished

technique. Out of this frustration came the popular joke that PCR should stand for

’pipette, cry, repeat’. To ensure that this stays a joke from now on, and that your PCR reactions

never drive you to despair again, we have compiled the most important tips and tricks for a

successful PCR set-up.

What is PCR?

The polymerase chain reaction (PCR) is used to amplify specific DNA sequences for

downstream use. Its inventor Kary Mullis, whose patent on PCR was approved in 1987, was

awarded the Nobel Prize in Chemistry six years later,1 and since this time, PCR has remained

one of the most essential molecular biology techniques. Genetic engineering, genotyping,

sequencing and the identification of familial relationships, to name a few examples, wouldn't be

possible without it.

PCR tips and tricks

To perform PCR reactions, you need to prepare a master mix, add template DNA, and amplify

the sequence of interest using a thermal cycler. This might seem straightforward, but it is far

from it. Calculating the required amounts of master mix reagents correctly to get the right

volume, at the right concentration, is the first challenge.

Once this is accomplished, the reagents need to be mixed together. The difficulty here is that

the liquids usually have to be cooled and they are often highly viscous, sticky and needed

in minimal quantities. In addition, work must be performed in a concentrated manner, as

distractions or interruptions can quickly lead to a situation where you no longer know which

reagents have already been added to the master mix. Errors such as skipping a tube or well can

CHAPTER 1: What you need to know about PCR

Simple PCR tips that can make or break your success

16

also easily occur when filling PCR strips or plates with master mix and adding template DNA,

especially when using single channel pipettes.

The last and probably biggest challenge is to keep your PCR reactions free from contamination.

PCR is a very sensitive assay that can create a large number of nucleic acid copies from a tiny

amount of starting material, so amplicon and sample contamination can be a huge problem.

Master mix calculations

Let's first have a look at the mathematical calculations needed to set up a PCR master mix.

We'll assume that you want to set up several PCR reactions with a volume of 50 μl each.

To calculate the required volume for each reagent, it is best to create a table (see Table 1) with

the necessary components, and fill in the stock concentrations and desired final concentrations

for the buffer, the MgCl2, the dNTPs and the primers. Then, calculate the dilution factors by

dividing the stock concentration by the final concentration. To determine the volume needed for

a single PCR reaction, divide the desired reaction volume by the dilution factor.2

For the polymerase, a slightly different equation is needed. The manufacturer of the enzyme

will tell you the amount of polymerase in one μl, e.g. 5 Units/μl. Fill in this value in the column

for the stock concentration and put the desired amount – e.g. 1.25 Units – in the column for

the final concentration. The volume needed can then be calculated as follows: 1.25 Units x

(1 μl / 5 Units) = 0.25 μl.3

The template DNA volume required depends on your sample type. You should add about 1 pg

to 10 ng of plasmid or viral DNA, and 1 ng to 1 μg of genomic DNA. In the example below, we

calculated how much you would need to use for 0.5 μl of a 1 μg/μl template DNA.4

Finally, add the required volumes for all the reagents. The difference between the desired total

reaction volume (50 μl) and the result obtained gives you the volume of PCR-grade water.5

REAGENT STOCK CONC. FINAL CONC.

(CF)

DILUTION

FACTOR

(= STOCK

CON. / CF)

VOLUME NEEDED

(= 50 ΜL / DIL.

FACTOR)

Buffer 10X 1X 10 5 μl

MgCl2 25 mM 1.5 mM 16.66 3 μl

dNTPs 10 mM 0.2 mM 50 1 μl

Forward primer 10 μM 250 nM 40 1.25 μl

Reverse primer 10 μM 250 nM 40 1.25 μl

Polymerase 5 Units/μl 1.25 Units - 0.25 μl

Template DNA 1 μg/μl - - 0.5 μl

PCR-grade water - - - 37.75 μl

Table 1: Example of a PCR master mix table

CHAPTER 1: What you need to know about PCR

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After determining the required reagent volumes for one PCR reaction, you can simply multiply

them by your sample number (plus the negative and positive controls) to get the total volumes

for the entire PCR set-up. We recommend adding one additional aliquot to that result, as some

of the master mix may be lost during pipetting due to evaporation, adherence to the tip, or

pipetting inaccuracies.

That's it, you are now ready to set up your PCR reactions by following the best pipetting

practices listed below.

Best PCR pipetting practices

Start by preparing your master mix from all the components listed above, except the template

DNA. The huge advantage of preparing the entire quantity of master mix needed for an

experiment, and subsequently transferring single aliquots into PCR strips or plates, is that

you can pipette higher volumes with better accuracy. On top of that, it reduces pipetting steps,

making the entire process less tiring and error prone. Since pipetting mistakes cannot be

completely ruled out, you should add the master mix components in order of their price, starting

with the most affordable reagent. This way, you waste less money if you have to start over.6

Once your master mix is finished, well mixed and dispensed into tubes or plates, you can

add the template DNA. As the DNA samples are usually highly viscous and needed in small

quantities, you should either dispense them into the master mix or onto the wall of the tube or

well. After dispensing, keep the plunger depressed while dragging the tip gently along the wall

of your labware to remove any residual liquid. In addition, we recommend using low retention

tips.

If you're not using a hot start polymerase, cool your reagents throughout the entire process of

master mix preparation and sample addition, to prevent non-specific amplification.

When you are ready to load your samples into the thermal cycler, check that they are tightly

capped or sealed, and spin them down to ensure that no droplets remain on the labware wall

during amplification.

Pipetting solutions for PCR reactions

Before discussing various pipetting solutions, we would like to address one of the most

important aspects of liquid handling. No matter which pipettes you choose, ensure that they are

well maintained by regularly calibrating them and checking their performance in between uses.

The most affordable pipettes for master mix preparation would be manual single channel

models. However, as you need to accurately measure and mix several very expensive

reagents, we recommend investing in electronic single channel pipettes. The motor-controlled

piston movement guarantees that they always dispense the exact desired volume, minimizing

variability to increase the precision and accuracy of pipetting.

For the container, you can either prepare the master mix in a tube or, if you intend to transfer

it with an electronic multichannel pipette, in a low dead volume reagent reservoir. The

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ASSIST PLUS pipetting robot transferring master mix into a 384 well PCR plate

combination of an electronic multichannel pipette and a reservoir is ideal for this step, because

you can fill several tubes or wells simultaneously. On top of that, electronic multichannel

pipettes usually feature a Repeat Dispense mode, allowing you to aspirate a large volume

of master mix, then dispense it into multiple smaller aliquots. It is also possible to use an

electronic single channel pipette if you have a low throughput.

To add template DNA to the master mix aliquots, an adjustable tip spacing pipette can be

very handy if the labware format of your samples doesn't match the container used for PCR

amplification. For example, it allows you to transfer several template DNA samples from

microcentrifuge tubes to an entire row or column of a 96 well plate in one step.

High throughput labs might even want to take advantage of automated solutions for master

mix plating and sample transfer, such as a pipetting platform that is capable of automating

electronic pipettes.

CHAPTER 1: What you need to know about PCR

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How to prevent PCR contamination

Several preventative measures should be taken to avoid contaminating your master mix or

template DNA with amplicons that were generated during previous PCRs.

One of the most effective means is to physically separate the master mix preparation, template

DNA addition, amplification and analysis areas from one another, and to work in laminar flow

or biosafety cabinets. Each work zone, and its corresponding equipment, should be cleaned

before and after an experiment, and tools used in one area should never enter another one.

When it comes to consumables, make sure you purchase sterile products that are certified to

be free from DNase, RNase and PCR inhibitors. Pipette tips should form a perfect seal with

the pipette to eliminate contamination that may occur when tips drip or fall off. Using filter tips

will also avoid the risk of aerosols entering your pipettes and contaminating subsequent PCR

reactions.

As you're a potential source of contamination too, always wear gloves to prevent introducing

enzymes, microbes and skin cells to the reaction, and change them when going from one area

to another. On top of that, keep your tubes closed whenever possible during the entire PCR

set-up.

Despite these preventative measures, you can't completely eliminate the possibility of

contaminated PCR reactions. To avoid having to throw away your entire stock of a certain

reagent if this occurs, prepare single use aliquots of your master mix components. You can also

prepare aliquots of positive and negative controls, as well as serial dilutions of standards for

quantitative PCR (qPCR) assays, ahead of time. Electronic pipettes with repeat dispense and

serial dilute modes can be helpful for this task, not only to reduce the risk of contamination, but

also to increase the efficiency of PCR set-up.

Conclusion

PCR is a fundamental technique in research, diagnostics and forensics. It often involves

pipetting minuscule reagent volumes with tricky properties, so it can be difficult to obtain the

desired results. On top of that, contamination can have a huge impact on results, as it's a very

sensitive assay. We hope that the tips and tricks provided in this article will help you make

your future PCR reactions a success. Many of these recommendations can also be applied to

other amplification assays, such as reverse transcription and qPCR, loop-mediated isothermal

amplification (LAMP) and helicase-dependent amplification (HDA).

CHAPTER 1: What you need to know about PCR

20

1.3 Setting up a PCR lab from scratch

PCR reactions are very sensitive and create a large number of copies of nucleic acids

from minute amounts of starting material. This makes them a fundamental and highly

effective molecular biology technique. However, because it is prone to amplicon and sample

contamination, planning and designing of your PCR lab space will need careful consideration.

CHAPTER 1: What you need to know about PCR

Designing your PCR lab

Ideally, a PCR lab should have two rooms with two areas, each designed for specific tasks.

The first room should be exclusively used for pre-PCR activities and divided into a master mix

preparation area and a sample preparation area. Air pressure should be slightly positive to

prevent aerosols from flowing in.

The second room should have a dedicated area for nucleic acid amplification, and another one

for product analysis. Air pressure should be slightly negative to ensure that amplicon aerosols

don't leave the room.

If you're lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible.

Having pre-PCR activities spatially separated from the amplification and analysis area – either

in different rooms or in separate benches – is very important, because you usually have a

low amount of the nucleic acid sample during preparation and a very high concentration after

CHAPTER 1: What you need to know about PCR 21

amplification. This means that if you analyze your PCR in the same space as you prepare your

master mix and samples, you may get false-positive results due to amplicon contamination.

You should also ensure that your lab set-up follows a unidirectional workflow. No materials or

reagents used in the amplification and analysis areas should ever be taken into the pre-PCR

space without a thorough decontamination. This means that you'll need dedicated equipment

for each area, e.g., two different sets of pipettes. This unidirectional workflow should also apply

to lab staff. If you've been working in the amplification and analysis areas, and you need to go

back to the pre-PCR area, change your personal protective equipment, as it may have been

contaminated by amplicon aerosols.

Another precautionary measure to take into account when setting up your PCR lab, in addition

to the spatial separation, is temporal separation. You could, for example, consider setting up

your PCR reactions in the morning, and perform the amplification and analysis steps in the

afternoon. This may limit your flexibility, but will prevent contamination issues and having to

repeat your experiment.

PCR equipment tips

PCR labs typically require a variety of equipment, such as centrifuges, vortex mixers, pipettes,

fridges and freezers, thermal cyclers and analysis instruments (e.g., electrophoresis systems).

Depending on the size of your lab and your applications, the amount of equipment you’ll need

may vary. Instead of providing you a 'shopping list', we will outline what you should look for

when purchasing equipment and consumables in order to keep contamination of your PCR

reactions to a minimum.

22 CHAPTER 1: What you need to know about PCR

Laminar flow or biosafety cabinet

Since you can never be 100 % certain that there are no amplicon aerosols in your pre-PCR

space, you should set up your PCR reactions in a laminar flow hood or biosafety cabinet,

decontaminated with a bleach solution prior to starting and after you finish your work.

Pipette tips and other consumables

Despite being more expensive than normal pipette tips, using filter tips for your PCR set-up will

avoid aerosols entering and contaminating your pipette, and avoid aerosols that might already

be present in your pipette contaminating your master mix or samples. To minimize your filter tip

consumption, first fill all your tubes with the master mix using only one tip or set of tips – if you're

using multichannel pipettes – and follow with your samples, using one tip per sample. Adding

the sample last is also recommended because it's easier to dispense it into a liquid than into an

empty tube, and because it reduces the risk of aerosolizing your sample as you pipette.

For consumables, you should make sure that you have enough small vials available in your lab

when your PCR reagents arrive. Aliquoting them into smaller containers will increase their shelf

life and prevent them from going through too many freeze/thaw rounds. If your reagents get

contaminated, it will also save you from throwing away your entire supply, as you’ll have clean

aliquots available for a second PCR.

Finally, you’ll need to make sure that all consumables and equipment are free of DNase, RNase

and PCR inhibitors. Always choose sterile products from manufacturers that can certify that

their tips and consumables are free of any of these potential contaminants.

CHAPTER 1: What you need to know about PCR 23

Cleaning and contamination control

You won’t need to worry about cleaning or contamination control when setting up your lab, but

you will when your lab is up and running. We will briefly address this topic below.

Whether you decide to set up your PCR reactions in a laminar flow hood, a biosafety cabinet

or an open bench, you will need to decontaminate your work space before and after set-up by

wiping it with a freshly made bleach solution and distilled water. The same process should be

performed in the amplification and analysis areas. You should also make sure you clean your

pipettes, equipment, doorknobs, and the handles of your fridges and freezers regularly.

Because PCR assays are so sensitive, all the preventative measures described here may still

not guarantee that your experiments will never get contaminated. It is therefore necessary

to include the appropriate controls to detect contamination early. Always include negative

and positive controls, as this will help identifying master mix contaminations, and confirm the

performance of the extraction protocol, reagents and amplification steps. Additionally, you

should monitor the positivity rate in your lab, and ensure that unexpected increases in detection

have identifiable causes, e.g., a seasonal outbreak.

Conclusion

In this article, we covered how to set up your PCR lab to ensure spatial and temporal

separation, and prevent contamination. We also outlined the key factors to consider when

purchasing equipment and consumables for your lab, to maintain safety and reduce wastage.

Lastly, we highlighted the importance of regular workspace cleaning and the use of appropriate

controls to detect any contamination early on. We hope that you are still just as excited about

setting up your PCR lab, and that this article has made the task less daunting for you.

24 CHAPTER 1: What you need to know about PCR

1.4 qPCR: How SYBR® Green and TaqMan®

real-time PCR assays work

qPCR, or real-time PCR, is a widely used method to quantify DNA sequences in samples.

This article gives you a comprehensive introduction to the topic, explaining how dye-based

and probe-based qPCR assays (like SYBR Green and TaqMan) work, how to validate your

amplification experiments, and how to analyze your qPCR data.

qPCR vs PCR vs RT-PCR

Before explaining how qPCR works, we would like to briefly outline its difference from standard

PCR and RT-PCR.

Whereas standard PCR monitors DNA amplification upon reaction completion, qPCR uses

fluorescent signals to monitor DNA amplification as the reaction progresses. This is why qPCR

is also referred to as real-time PCR, quantitative PCR or quantitative real-time PCR.

RT-PCR, not to be confused with real-time PCR, stands for reverse transcription PCR and can

be used to amplify RNA target sequences. It involves an initial incubation of the sample RNA

with a reverse transcriptase enzyme and a DNA primer before amplification.

How qPCR works

qPCR relies on fluorescence from intercalating dyes or hydrolysis probes to measure DNA

amplification after each thermal cycle. The most common dye-based method is SYBR Green,

and the most common probe-based method is TaqMan, which is why this article will focus on

these two qPCR techniques.

CHAPTER 1: What you need to know about PCR 25

SYBR Green qPCR

Like standard PCR, the SYBR Green protocol consists of denaturation, annealing and

extension phases. The difference being that you add some double-stranded DNA binding dye,

SYBR Green I, to your master mix during qPCR setup. This fluorescent dye intercalates into

double-stranded DNA sequences during the extension phase, where it shows a strong increase

in fluorescent signal. Measuring this signal at the end of every thermal cycle will allow you to

determine the quantity of double-stranded DNA present.

The downside of the SYBR Green assay is that the dye binds to any double-stranded DNA

sequence. This means that you could also detect fluorescence emitted from non-specific

qPCR products, such as primer dimers. To eliminate this risk, check the reaction specificity by

performing a melting curve analysis, explained later in the article, or use the TaqMan method.

TaqMan qPCR

Instead of using intercalating dyes, this assay uses TaqMan probes with a 5' fluorescent

reporter dye and a 3' quencher dye. These probes are target-specific, and only bind to the

DNA sequence of interest downstream of one of the primers during the annealing step. When

the enzyme Taq-polymerase encounters the TaqMan probe during the extension phase, it

displaces and cleaves the 5' reporter dye. Once the reporter dye has been separated from

the quencher dye, its measurable fluorescent signal at the end of every qPCR cycle increases

significantly. The second DNA strand is synthesized in parallel but, as no probe is attached to it,

this process can't be monitored by fluorescence measurements.

Compared to the SYBR Green assay, the use of TaqMan probes is more expensive, but also

offers two significant advantages:

• the TaqMan assay only measures amplification progression of the target sequence, as the

probes are target specific.

• you can monitor the quantity of various qPCR products in a single reaction by adding different

primers and TaqMan probes with different reporter dyes to the master mix. This multiplex

approach allows you to detect several fluorescent signals at the end of every thermal cycle.

26 CHAPTER 1: What you need to know about PCR

Amplification plot

For both qPCR methods, data is visualized in an amplification plot, with the number of thermal

cycles on the x-axis, and the fluorescent signals detected on the y-axis:

CHAPTER 1: What you need to know about PCR 27

As can be seen, fluorescence remains at background levels during the first thermal cycles.

Eventually, the fluorescent signal reaches the fluorescence threshold, where it is detectable over

the background fluorescence. The cycle number at which this happens is called the threshold

cycle (Ct). If the Ct value for a sample is high, it means that little starting material was present,

and vice versa. Please note that you should always analyze at least three replicates of each

sample, as tiny pipetting errors during qPCR set-up can result in huge differences in Ct values.

The Ct value is sometimes also referred to as crossing point (Cp), take-off point (TOP) or

quantification cycle (Cq) value, with MIQE guidelines suggesting using Cp value to standardize

terminology.1 In this article we will continue to call it Ct, as this is the most commonly used term.

MIQE guidelines

The MIQE (Minimum Information for Publication of Quantitative Real-Time PCR Experiments)

guidelines describe the minimum information necessary for evaluating qPCR experiments.

When publishing a manuscript, the scientist needs to provide all relevant experimental

conditions and assay characteristics described by the MIQE guidelines, allowing reviewers

to assess the validity of the protocols used, and enabling other scientists to reproduce the

experiments.

Validation of qPCR assays

qPCR amplification plots can be analyzed using absolute or relative quantification. However,

before explaining qPCR data analysis, we need to quickly discuss how to determine reaction

efficiency and specificity. You don't need to perform these steps after every qPCR experiment,

but should always validate these two values when setting up a new qPCR protocol or changing

your current workflows.

Reaction efficiency

A perfect qPCR assay would have a reaction efficiency of 100 %, which means that the number

of template DNA copies would double at every thermal cycle. As this is almost impossible to

achieve in practice, reaction efficiencies between 90 and 110 % are considered to be ideal.

To calculate the reaction efficiency of your assay, you need to set up a 10-fold serial dilution

of an undiluted sample with a known amount of template DNA. After running a qPCR, create a

standard curve with the log of the starting quantity on the x-axis and the Ct values on the y-axis.

28

Using the equation for the linear regression line (y = mx + b), you can now determine the

reaction efficiency as follows2:

Efficiency = (10(-1/m)-1) x 100

In our example, m would be -3.5826, resulting in a reaction efficiency of 90.1634 %.

Reaction specificity

Reaction specificity can be determined using a melting curve analysis, allowing you to identify

non-specific qPCR products and primer-dimers. To perform a melting curve analysis, run a

qPCR assay with a fluorescent intercalating dye like SYBR Green I. After amplification, the

thermal cycler increases the temperature step by step while monitoring fluorescence. As

the temperature increases, the dsDNA qPCR products present will denature, resulting in a

decreasing fluorescent signal:

CHAPTER 1: What you need to know about PCR

CHAPTER 1: What you need to know about PCR 29

Then, plot the change in slope of this curve as a function of temperature to obtain a melting

curve:

If you're observing only one melting peak like the image above, your qPCR assay is specific.

If there are several melting peaks, primer-dimers and/or non-specific products were amplified

during qPCR, and you should redesign your experiment to increase its specificity.

30 CHAPTER 1: What you need to know about PCR

Analysis of qPCR data

qPCR data can be analyzed by absolute or relative quantification, and the method suitable

for your experiment depends on your goal. Absolute quantification allows you to determine

the quantity of starting material that was present in a given sample before amplification. For

example, this method can be used to determine the viral load of a patient sample. Relative

quantification is applied to compare levels or changes in gene expression between different

samples. For example, it is helpful to investigate whether the expression of a certain gene is

higher in a tumor sample than in a healthy control sample.

Absolute quantification

After qPCR amplification, you will have produced an amplification plot, and know the Ct value

of each sample. To find the quantity of starting material present in your samples, you need to

compare these values to a standard curve. As seen above in the section on reaction efficiency,

a standard curve is obtained by amplifying a serial dilution of a sample with a known amount of

template DNA, then plotting the Ct values against the log of the starting quantities.

The equation for the linear regression line of the standard curve (y = mx + b) will then allow you

to calculate the quantity of starting material for each sample. As y corresponds to the Ct value,

and x to the log quantity, the equation for the linear regression line is equivalent to:

Ct = m(log quantity) + b

Solving this equation for the quantity will give you the formula:

Quantity = 10((Ct-b)/m)

This will allow you to quickly determine the quantity of starting material in each sample.

Y = mx + b → Ct = m(log quantity) + b → Quantity = 10((Ct-b)/m)

Relative quantification

To compare levels or changes in target gene expression between different samples and a

control sample, you first need to define a reference gene whose expression is unregulated.

Then, run a qPCR to obtain the Ct values for the reference gene, target gene in your samples,

and the control sample.

If the reaction or primer efficiencies for the reference and target genes are near 100 %, and

within 5 % of each other, you can then use the ΔΔCt method – also called the Livak method – to

determine the expression rate of the target gene in your samples. However, if the efficiencies

are further apart, you should use the Pfaffl method. To learn how to calculate reaction

efficiencies, please refer to the 'Reaction efficiency' section earlier in the article.

CHAPTER 1: What you need to know about PCR 31

The calculations for the two methods are as follows:

ΔΔCt method

Normalize the Ct of the target gene to the Ct of the reference gene for each sample and the

control sample:

ΔCt(sample) = Ct(target gene) – Ct(reference gene)

ΔCt(control) = Ct(target gene) – Ct(reference gene)

Normalize the ΔCt of each sample to the ΔCt of the control sample:

ΔΔCt(sample) = ΔCt(sample) – ΔCt(control)

Since the calculations are in logarithm base 2, you must use the following equation to get the

normalized expression ratio for each sample:

Normalized expression ratio = 2-ΔΔCt(sample)

Pfaffl method

Calculate the ΔCt of the target gene for each sample:

ΔCt(target gene) = Ct(target gene in control) – Ct(target gene in sample)

Calculate the ΔCt of the reference gene for each sample:

ΔCt(reference gene) = Ct(reference gene in control) – Ct(reference gene in sample)

Calculate the normalized expression ratio for each sample:

Normalized expression ratio = ((Etarget gene)ΔCt(target gene)) / ((Ereference gene)ΔCt(reference gene))

Etarget gene: Reaction efficiency of the target gene

Ereference gene: Reaction efficiency of the reference gene

The normalized expression ratio obtained using the ΔΔCt or the Pfaffl method is the fold

change of the target gene in your sample relative to the control. A normalized expression ratio

of 1.2 would mean that you have a gene expression of 120 % relative to the control.

Conclusion

We hope that this article answered all your questions regarding qPCR methods, assay

validation and data analysis.

32

1.5 How to design primers for PCR

PCR is one of the most widespread molecular biology applications, yet it is anything but simple

to perform. Common issues – such as a low product yield or non-specific amplification – are

often caused by poorly designed PCR primers. We have therefore summarized the most

important information on designing PCR primers to help you overcome these challenges.

What is a PCR primer?

Primers – also called oligonucleotides or oligos – are short, single-stranded nucleic acids used

in the initiation of DNA synthesis. During PCR reactions, they anneal to the plus and minus

strands of the template DNA, flanking the sequence that needs to be amplified.

How to design PCR primers?

PCR primers have to be tailored to both the region of interest of your template DNA and your

reaction conditions. This means that, unlike the other components of the PCR master mix, you

can't just buy them, but need to design them yourself using a primer design tool. These tools

allow you to set parameters such as primer length, melting temperature, GC content and more.

Read on to learn what the optimal values for each of these parameters are, and how they affect

your PCR assay.

CHAPTER 1: What you need to know about PCR

CHAPTER 1: What you need to know about PCR 33

Primer length

The optimal length of a PCR primer lies between 18 and 24 bp. Longer primers are less efficient

during the annealing step, resulting in a lower amount of PCR product. Conversely, shorter

primers are less specific during the annealing phase, leading to more non-specific binding and

amplification. However, there are exceptions to this rule. For example, some scientists have

successfully used miniprimers that are 10 bp long to expand the scope of detectable sequences

in microbial ecology assays.

Target sequence length

The target sequence to be amplified should ideally be between 100 and 3000 bp for standard

PCR assays, and 75 and 150 bp for qPCR assays. Longer sequences usually need special

enzymes and reaction conditions to ensure that they are completely and specifically amplified.

Primer melting temperature

The primer melting temperature (Tm) can be defined as the temperature at which half of the

primers dissociate from the template DNA. It is usually between 50 and 60 °C, and the melting

temperatures of the forward and reverse primers should be within 5 °C of each other. If the two

melting temperatures are further apart, it won't be possible to find an annealing temperature

that allows both primers to bind to the template DNA.

Most primer design tools use the nearest neighbor method to calculate primer melting

temperatures, as it's the most accurate. However, if you want to make an approximate

calculation yourself, you can use this formula:

Tm = 4 °C x (G+C) + 2 °C x (A+T)

Tm: melting temperature

G, C, A, T: number of nucleobases (guanine, cytosine, adenine, thymine) in the primer

As indicated in the formula above, G-C bonds are harder to break than A-T bonds – because

G-C base pairs are linked by three hydrogen bonds, and A-T base pairs by two – and the length

of the primer also impacts its melting temperature. This means that you can either increase the

GC content of a primer (provided the template allows for this), or slightly extend its length if its

melting temperature is too low.

34

Primer annealing temperature

The primer annealing temperature (Ta) is the temperature needed for the annealing step of

the PCR reaction to allow the primers to bind to the template DNA. The theoretical annealing

temperature can be calculated as follows:

Ta = 0.3 x Tm(primer) + 0.7 x Tm(product) – 14.9

Ta: primer annealing temperature

Tm(primer): lower melting temperature of the primer pair

Tm(product): melting temperature of the PCR product

Once you've calculated the theoretical annealing temperature, the optimal annealing

temperature needs to be determined empirically. To achieve this, perform a gradient PCR,

starting a few degrees below the calculated annealing temperature, and ending a few degrees

above. After amplification, run a gel, and the sample producing the clearest band contains the

largest quantity of PCR product, making its annealing temperature the optimal one for your

primers. Usually, you'll get a value that is 5 to 10 °C lower than the primer melting temperature.

CHAPTER 1: What you need to know about PCR

35

It's important to determine the optimal annealing temperature, as primers could form hairpins or

bind to regions outside the DNA sequence of interest if it's too low, producing non-specific and

inaccurate PCR products. If the annealing temperature is too high, the primers won't sufficiently

bind to the template DNA, and you'll obtain little to zero amplicons.

CHAPTER 1: What you need to know about PCR

GC content

As seen before, G-C base pairs are stronger than A-T base pairs, which means that a higher

GC content ensures a more stable binding between the primers and the template DNA. The

optimal GC content of a primer lies between 40 and 60 %, and primers should have two to three

Gs and Cs at the 3' end to bind more specifically to the template DNA.

Runs and repeats

Avoid runs of four or more single bases – such as ACCCCC – or dinucleotide repeats (for

example, ATATATATAT), as they can cause mispriming.

Cross homology

If a primer is homologous to a template DNA sequence outside the region of interest, these

sequences will be amplified too. Therefore, you should always test the specificity of your

primer design against genetic databases; for example, by ‘blasting’ them through NCBI BLAST

software.

36

Your PCR product yield will be less if secondary structures form and remain stable above the

annealing temperature of your reaction, as the primers bind to themselves or another primer

instead of the template DNA. This is why your primer design tool should be able to check for,

and warn you of stable secondary structures.

Mismatches and degenerated positions

Mismatches are primer bases that aren't complementary to the target sequence. They can be

tolerated to a certain extent, and are sometimes even necessary; for example, when performing

a multi-template PCR to amplify a set of similar target sequences from different bacteria with

a single set of primers. Degenerate primers could help if mismatches negatively impact the

performance of your PCR.

Degenerate primers have several different nucleotides in some of their positions. For example,

instead of A you could have an equal concentration of A and T in a certain position. The codes

for the different nucleotide combinations available for degenerate primers are as follows:

CHAPTER 1: What you need to know about PCR

Secondary structures

There are three different types of secondary structures – also called primer dimers – that can

form during a PCR assay:

• Hairpins: caused by intra-primer homology – when a region of three or more bases is

complementary to another region within the same primer – or when a primer melting

temperature is lower than the annealing temperature of the reaction.

• Self-dimers: formed when two same sense primers have complementary sequences – interprimer

homology – and anneal to each other.

• Cross-dimers: formed when forward and reverse primers anneal to each other when there is

inter-primer homology.

37

IUPAC NUCLEOTIDE CODE BASE

R A or G

Y C or T

S G or C

W A or T

K G or T

M A or C

B C or G or T

D A or G or T

H A or C or T

V A or C or G

N Any base

Conclusion

This article summarized the key points to consider when designing PCR primers to help avoid

common issues like low product yield or non-specific amplification. We covered optimal primer

and target sequence lengths, and ideal primer melting and annealing temperatures. We also

provided helpful tips for other crucial factors such as GC content, runs and repeats, cross

homology and the danger of stable secondary structures. Lastly, the article highlighted the

value and pitfalls of mismatches and degenerated positions. That's it, after reading about all of

this, you are sure to be a 'PCR Primer Pro'!

CHAPTER 1: What you need to know about PCR

38

CHAPTER 2:

INTEGRA Biosciences’ PCR solutions

PCR is a robust method, but it’s comprised of numerous stages, involving multiple precise

pipetting steps that often prove time consuming and prone to errors. Temperature-sensitive

reagents and samples may affect accuracy, and the varying viscosities of samples, as well as

‘sticky’ DNA, can be difficult to handle. On top of this, the repetitive nature of this work can also

frequently result in user fatigue and handling mistakes.

Fortunately, the right tools can eliminate your pipetting predicaments, vastly improving the

reproducibility and productivity of your PCR workflows. Here, we will demonstrate how our

range of liquid handling solutions are perfect for PCR applications, allowing you to create a

faster and more efficient workflow with fewer errors.

Manual and electronic pipettes

A good starting point for lower throughput PCR applications – up to half a plate per day – is our

EVOLVE single or multichannel pipettes, which feature convenient volume adjustment dials to

increase the accuracy and speed of manual handling. Our range of VIAFLO electronic pipettes

is also suitable for low throughput PCR set-up, and can easily handle up to eight plates per day.

CHAPTER 2: INTEGRA Biosciences’ PCR solutions

Learn more

about

EVOLVE

Learn more

about

VIAFLO

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 39

Learn more

about

VOYAGER

Adjustable tip spacing pipettes

PCR set-up usually requires transferring liquids between different labware formats which is

tedious and highly error prone. Our VOYAGER adjustable tip spacing pipettes solve these

problems, increasing speed and eliminating transfer errors, while ergonomic single-handed

operation leaves the other hand free to handle labware.

96 and 384 channel pipettes

We have a wide range of options perfect for productive high throughput PCR set-up – more

than eight plates per day – which are suitable for different lab sizes and budgets. Our

VIAFLO 96 and VIAFLO 384 channel handheld electronic pipettes, as well as the

MINI 96 channel portable electronic pipette, can reduce handling steps while

increasing productivity and reproducibility.

Learn more

about

MINI 96

Learn more

about

VIAFLO 96

and VIAFLO 384

40 CHAPTER 2: INTEGRA Biosciences’ PCR solutions

Pipetting robots

INTEGRA also offers pipetting robots for high throughput laboratories, or for labs that want to

reduce the risk of contamination due to manual processing. For example, the ASSIST PLUS

pipetting robot can automate the D-ONE single channel pipetting module for master mix

preparation, and VIAFLO and VOYAGER multichannel pipettes to take care of the multiple

pipetting steps in PCR workflows.

Learn more

about

D-ONE

Learn more

about

GRIPTIPS

Learn more

about

ASSIST PLUS

Pipette tips

INTEGRA has developed GRIPTIPS pipette tips to complement its range of pipetting solutions.

GRIPTIPS are free from RNase, DNase and PCR inhibitors, and perfectly fit all INTEGRA

pipetting solutions, reducing the risk of contamination from tips that leak or fall off.

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 41

Learn more about

sample transfers

from plate to plate

Learn more about

sample transfers

from tubes to plates

Sample reformatting

The transfer of samples between different labware formats is a slow, tedious and highly

error-prone procedure when performed manually with a single channel pipette. The

combination of the ASSIST PLUS pipetting robot and VOYAGER adjustable tip spacing

pipette provide a novel solution for automated, accurate and efficient liquid transfer of multiple

samples in parallel. For even higher throughput applications, the VIAFLO 96, VIAFLO 384

and MINI 96 offer a fast solution for whole plate transfers.

42 CHAPTER 3: Application Notes

CHAPTER 3:

Application notes

Our pipetting instruments are used across a broad spectrum of life sciences applications. To

help share this knowledge and experience of using INTEGRA products with the wider scientific

community, we have developed an application database which contains a wide range of useful

application notes. Here are some of the most relevant app notes related to PCR protocols and

workflows.

3.1 Efficient and automated 384 well qPCR set-up

with the ASSIST PLUS pipetting robot

Using the ASSIST PLUS pipetting robot to automate set-up for a 384 well

plate qPCR

Setting up a qPCR is a tedious process consisting of multiple pipetting steps. One particularly

challenging task is reformatting from microcentrifuge tubes into a 384 well plate, which is time

consuming and requires a lot of concentration. Another common problem is the loss of valuable

and expensive substances, such as master mix and

precious samples, due to the reservoir dead volume.

The ASSIST PLUS pipetting robot, in combination with

the VIAFLO and VOYAGER electronic pipettes,

streamlines the workflow and increases the throughput

and the reproducibility of qPCR set-ups, with minimal

manual input. The loss of expensive substances or

valuable samples due to reformatting errors is

eliminated. The unique design of the ASSIST PLUS

pipetting robot, together with the intuitive

VIALAB software, offers

exceptional flexibility and

straightforward implementation.

CHAPTER 3: Application Notes 43

Key benefits

• Automating the qPCR set-up with the

VIAFLO 16 channel electronic pipette and

the ASSIST PLUS pipetting robot allows

considerably faster sample preparation,

freeing up time for scientists to focus on

other experiments.

• Automation of VOYAGER adjustable tip

spacing pipettes with the ASSIST PLUS

offers a reliable pipetting method that

requires minimal manual intervention and

eliminates the risk of reformatting errors.

• The use of low retention GRIPTIPS with

heightened hydrophobic properties and

SureFlo™ low dead volume reservoirs with

an anti-sealing array helps to save precious

samples and master mix. Combined with

the high pipetting accuracy and precision

of the ASSIST PLUS pipetting robot, this

enables exceptionally low dead volumes to

be achieved.

• The ASSIST PLUS pipetting robot, in

combination with the intuitive VIALAB

software, is quick to set up and easy to use.

Overview: qPCR set-up

The ASSIST PLUS pipetting robot is used to set up a 384 well format qPCR by pipetting 64

samples in triplicate with two different master mixes for the detection of two genes of interest

(GOI 1 and GOI 2).

The protocol is divided into two programs that guide the user through all the steps of the qPCR

set-up:

• Program 1: Mastermix_qPCR

• Program 2: Samples_qPCR

The ASSIST PLUS pipetting robot operates a VIAFLO 16 channel 125 μl electronic pipette with

125 μl sterile, filter, low retention GRIPTIPS for program 1 and a VOYAGER 8 channel 12.5 μl

electronic pipette with 12.5 μl sterile, filter, low retention GRIPTIPS for program 2.

44

Experimental set-up: Program 1 - master mix

transfer (Mastermix_qPCR)

Prepare the pipetting robot deck as follows (Figure 1):

Deck position A: Dual reservoir adapter – 2 x 10 ml reagent

reservoir with SureFlo anti-sealing array

(Figure 2) containing master mix 1 and 2.

Deck position B: 384 well PCR plate, placed on an INTEGRA

cooling block in the landscape position.

CHAPTER 3: Application Notes

Figure 1: Set-up for the master mix transfer. Position A: dual reservoir adapter with 2 x 10 ml reagent

reservoirs with SureFlo anti-sealing array. Position B: 384 well PCR plate, placed on an INTEGRA

cooling block.

Figure 2: The INTEGRA dual reservoir adapter accommodates both 10 ml reagent reservoirs on one

deck position.

CHAPTER 3: Application Notes 45

Step-by-step procedure

1. Transfer master mixes into the 384 well plate

Add master mixes 1 and 2 into the left and right sides of the 384 well PCR

plate, respectively.

Use an EVOLVE 5000 μl manual pipette with

5000 μl sterile, filter, low retention GRIPTIPS to

fill the left 10 ml reagent reservoir with SureFlo

anti-sealing array with 1.6 ml of master mix 1 and

the right reservoir with 1.6 ml of master mix 2

(position A). Select and run the VIALAB program

‘Mastermix_qPCR’ on the VIAFLO 16 channel

125 μl electronic pipette with 125 μl sterile, filter,

low retention GRIPTIPS. The ASSIST PLUS

pipetting robot automatically transfers 7.5 μl of

master mix 1 (pink) into the left half of the 384 well

PCR plate and 7.5 μl of master mix 2 (blue) into the

right half (Figure 3) using the Repeat Dispense

mode with a tip touch on the surface of the liquid to

increase pipetting precision. Figure 4 shows the

pipetting robot transferring the master mix into a

384 well plate.

Tips:

• Pre- and post-dispense steps are recommended

to increase the accuracy and precision of

pipetting. The pre- and post-dispense volumes

should be between 3 and 5 % of the nominal

volume of the pipette.

• The low retention GRIPTIPS are made from a

unique polypropylene blend with heightened

hydrophobic properties for superior accuracy

and precision while pipetting viscous and low

surface tension liquids.

• The reservoirs’ SureFlo anti-sealing array and

a unique surface treatment that spreads liquid

evenly enable the pipette tips to sit on the bottom

and still aspirate liquids accurately, reducing

dead volumes.

Figure 3: Pipetting scheme for master mixes 1 (pink) and 2 (blue).

Figure 4: Example of the ASSIST PLUS pipetting robot

transferring a master mix into a 384 well PCR plate.

46 CHAPTER 3: Application Notes

Experimental set-up: Program 2 - sample transfer

(Samples_qPCR)

Prepare the pipetting robot deck as follows (Figure 5):

Deck position B: 384 well PCR plate, placed on an INTEGRA

cooling block.

Deck position C: INTEGRA 1.5 ml microcentrifuge tube rack,

with tubes containing samples 1-32.

Figure 5: Set-up for the sample transfer protocol. Position B: 384 well PCR plate, placed on an INTEGRA

cooling block. Position C: INTEGRA 1.5 ml microcentrifuge tube rack, with tubes containing samples 1-32

(Figure 6).

Figure 6: Example of the ASSIST PLUS pipetting samples from the INTEGRA microcentrifuge tube rack

into a 96 well plate.

VOYAGER - 12.5 μl – 8CH

12.5 μl GRIPTIP,

sterile, filter

B 384 well PCR Sapphire on 384 well cooling block – 45 μl C Rack for 1.5 ml microcentrifuge tubes – 1500 μl

CHAPTER 3: Application Notes 47

Step-by-step procedure

1. Sample transfer into the 384 well plate

Add the 64 samples in triplicate to the master mixes.

Place samples 1-32 in an INTEGRA 1.5 ml microcentrifuge tube rack on position C. Run the

VIALAB program ‘Samples_qPCR’ on a VOYAGER 8 channel 12.5 μl electronic pipette to start

the sample transfer. The ASSIST PLUS transfers 2.5 μl of the first 32 samples in triplicate into

master mixes 1 and 2 (Figure 7, yellow/brown), using the Repeat Dispense mode with a tip

touch on the side of the well to make sure that no droplets adhere to the GRIPTIPS. After this

step, a prompt informs the user to place the second series of samples (33-64) on position C.

The ASSIST PLUS pipetting robot continues by transferring 2.5 μl of the samples in triplicate

into the other half of master mixes 1 and 2 (Figure 7, green).

Tip: Use sterile, filter, low retention GRIPTIPS for optimal liquid recovery of precious solutions,

such as the master mix and samples.

Figure 7: Pipetting scheme of the qPCR assay.

master mix 1 master mix 2

Sample

33 - 64

Sample

1 - 32

48 CHAPTER 3: Application Notes

Remarks

VIALAB software:

The VIALAB program can easily be adapted to fit the user’s demands, especially if specific

labware, incubation times or protocols are needed.

Partial plates:

The programs can be adapted at any time to a different number of samples, giving

laboratories total flexibility to meet current and future demands.

Conclusion

• The time required for a 384 well qPCR set-up can be reduced from 1.5 hours using

a single channel pipette to 12 minutes using the ASSIST PLUS pipetting robot in

combination with VIAFLO 16 channel and VOYAGER 8 channel pipettes.

• The ASSIST PLUS, together with the VOYAGER adjustable tip spacing pipette,

guarantees perfectly reproducible test results and eliminates all risks of reformatting

errors when transferring samples from microcentrifuge tubes into a 384 well plate.

• INTEGRA’s low retention GRIPTIPS increase pipetting precision for viscous or low

surface tension liquids. The reagent reservoirs with SureFlo anti-sealing array reduce the

dead volume of costly reagents and precious samples.

• The intuitive VIALAB qPCR program is quick to set up and easy to use or adapt to other

pipetting protocols.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 3: Application Notes 49

3.2 Automated RT-PCR set-up for

COVID-19 testing

How to prepare RT-PCR plates for SARS-CoV-2 detection

with the ASSIST PLUS

The emergence and outbreak of the novel coronavirus

SARS-CoV-2 (COVID-19) has placed unprecedented

demands on laboratories testing for COVID-19, leaving

scientific staff to contend with a spiraling influx of patient

samples and a rapid, continuous growth in workload.

Laboratories need additional automated liquid handling

instruments for viral nucleic acid extraction

and RT-PCR set-up – which are the

most labor-intensive processes in

the diagnostic testing workflow – to

increase the sample throughput

capacity, reduce manual

intervention by laboratory

analysts and fast track the

development of COVID-19 assays.

The ASSIST PLUS pipetting robot together with a VOYAGER

adjustable tip spacing pipette, low retention GRIPTIPS and SureFlo

10 ml reagent reservoirs were successfully used for RT-PCR set-up in

COVID-19 testing laboratories.

50 CHAPTER 3: Application Notes

Key benefits

• The full automation capability of the

ASSIST PLUS reduces manual intervention

and frees highly valuable time for laboratory

personnel in this critical COVID-19

pandemic.

• The compact and easy-to-use

ASSIST PLUS pipetting robot allows

fast set-up regarding installation and

programming, allowing labs to immediately

increase their sample processing capacity

and fast track assay development for

COVID-19 sample testing.

• VOYAGER adjustable tip spacing pipettes

in combination with the ASSIST PLUS

provide unmatched pipetting ergonomics by

automatically reformatting patient samples

from tube racks into 384 well plates.

• Optimal pipette settings, including tip

immersion depth, pipetting speeds and

angles, deliver reproducible, precise and

accurate results, with no contamination

observed in controls or patient samples.

• The use of INTEGRA’s low dead volume,

SureFlo 10 ml reagent reservoirs, together

with low retention GRIPTIPS, demonstrated

excellent results, enabling efficient handling

of the precious and expensive one-step RTPCR

master mix used for patient testing.

Overview: Automated RT-PCR set-up

The ASSIST PLUS pipetting robot is used to automate testing of suspected COVID-19

positive cases in a 384 well plate. The pipetting robot operates a VOYAGER 12 channel 50 μl

electronic pipette with 125 μl sterile, filter, low retention GRIPTIPS. To double the available

testing capacity and, concurrently, decrease the cost per test of expensive one-step RT-PCR

reagents of dwindling availability, the total PCR reaction volume was miniaturized, reducing it

to 10 μl – inclusive of 7.5 μl one-step RT-PCR master mix and 2.5 μl of nucleic acid template.

The templates were extracted from combined nasopharyngeal/oropharyngeal flocked swabs

or sputum samples. The following procedure is based on the protocol used by the Microbiology

and Molecular Pathology Department at Sullivan Nicolaides Pathology (SNP) – part of the

Sonic Healthcare Group – in Brisbane, Australia.

The protocol is divided into two parts:

• Program 1: Add the master mix (1-COVID-19)

• Program 2: Add the nucleic acid template (2-COVID-19)

CHAPTER 3: Application Notes 51

Experimental set-up: Program 1

Deck position A: 10 ml reagent reservoir with

SureFlo anti-sealing array containing

3 ml of one-step RT-PCR master mix.

Deck position C: 384 well plate placed on a PCR 384 well

cooling block, allowing the master mix and

samples to be kept cold, and enabling exact

positioning of the PCR plate on the deck.

Figure 1: The set-up for program 1-COVID-19.

VOYAGER - 50 μl – 12CH

50/125 μl GRIPTIP, sterile, filter,

low retention

A Multichannel reservoir – 10ml C PCR cooling block 384_system

52 CHAPTER 3: Application Notes

Step-by-step procedure

1. Add the master mix

Fill the 384 well plate with the one-step RT-PCR master mix.

Place the one-step RT-PCR master mix in a 10 ml sterile, polystyrene reagent reservoir with

INTEGRA’s SureFlo anti-sealing array. Set up the deck with the required labware, as indicated

in Figure 1. Select the VIALAB program 1-COVID-19. The VOYAGER pipette automatically

transfers the master mix from the reservoir into the 384 well plate (LightCycler® 480 Multiwell

Plate, Roche) using the Repeat Dispense mode with tip touch. Each well of the plate is filled

with 7.5 μl of master mix.

Tips:

• Using a 10 ml reagent reservoir with SureFlo anti-sealing array allows a very low dead

volume (<20 μl) to minimize the loss of expensive reagent of dwindling availability

(see Figure 2).

• The combination of a low pipetting speed – set at 2 – and low retention GRIPTIPS shows

excellent results when pipetting the viscous and foamy master mix.

• Pre- and post-dispense settings, together with the tip touch option, guarantee reproducible,

precise and accurate pipetting results (see Figure 2).

• The PCR cooling block is used as a support to fix the position of the 384 well plate on the

deck, ensuring exact tip positioning when pipetting. The cooling block also helps to keep

samples and reagents cool if required by the protocol.

Figure 2: Precise and accurate dispensing of one-step RT-PCR master mix from the low dead

volume reagent reservoir to the 384 well plate.

CHAPTER 3: Application Notes 53

Experimental set-up: Program 2

Deck position A and B: FluidX Cluster 0.7 ml tubes containing the

nucleic acid templates. The tubes are stored

in a 96-format rack. A total of four sample

racks are used for the protocol (two on

position A and two on position B).

Deck position C: 384 well plate placed on a PCR 384 well

cooling block.

Figure 3: The set-up for program 2-COVID-19.

2. Add the nucleic acid templates

Transfer the samples from four 96-format tube racks to the 384 well plate.

Nucleic acid templates extracted from combined nasopharyngeal/oropharyngeal flocked

swabs or sputum samples are stored in FluidX Cluster 0.7 ml tubes placed in a 96-format

rack. The VOYAGER pipette transfers 2.5 μl of template from the tubes to the 384 well plate,

automatically changing the GRIPTIP pipette tips after each dispense. Both position A and B

are used to house the samples on the deck (see Figure 3). The pipette prompts the user when

it is time to replace the tube racks on the deck. After user confirmation, the VOYAGER pipette

continues reformatting the samples from tubes to the plate.

50/125 μl GRIPTIP, sterile, filter,

low retention

VOYAGER - 50 μl – 12CH

A FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl

B FluidX 96-formal, 0.7 ml Internal Thread Tube, V-Bottom

– 700 μl C PCR cooling block 384_system

54

Tips:

• The VOYAGER pipette’s tip spacing capability combined with automatic Tip Change ensures

easy and rapid sample transfer without risk of contamination or reformatting errors.

• Using an air gap of 1.5 μl when aspirating the viral nucleic acid template eliminates the risk of

contamination risk during pipette tip travel.

Note: Automated RT-PCR testing for COVID-19 with the ASSIST PLUS can also be

performed using a VOYAGER 8 channel 50 μl electronic pipette (see Figure 5).

Figure 4: Easy and rapid transfer of patient nucleic acid templates from the tube rack to the 384 well

plate using the VOYAGER adjustable tip spacing pipette together with the ASSIST PLUS pipetting robot.

Figure 5: Automated RT-PCR testing for COVID-19 using the ASSIST PLUS pipetting robot together with

a VOYAGER 8 channel adjustable tip spacing pipette, as performed in the Microbiology and Molecular

Pathology Department at SNP.

CHAPTER 3: Application Notes

55

Remarks

4 Position Portrait Deck:

If your process allows, the protocol can be compiled into one simple program using the

4 Position Portrait Deck option on the ASSIST PLUS (see Figure 6).

96 well plates:

The protocol can be readily adapted to 96 well format.

VIALAB software:

The VIALAB programs can be easily adapted to your specific labware and protocols.

CHAPTER 3: Application Notes

Figure 6: Example set-up of the 4 Position Portrait Deck when combining programs 1-COVID-19 and

2-COVID-19 in one program.

50/125 μl GRIPTIP, sterile, filter,

low retention

VOYAGER - 50 μl – 12CH

A Multichannel reservoir – 10ml B FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl

C FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl D PCR cooling block 384_system

56

Conclusion

• In the context of a global pandemic where laboratories are under increasing pressure to

analyze more and more patient specimens to confirm COVID-19 cases, testing labs can

rapidly benefit from the advantages of the ASSIST PLUS pipetting robot, allowing them to

increase their sample processing capacity.

• Pipetting results were reproducible, precise and accurate, with no contamination

observed in controls or patient samples.

• The ASSIST PLUS pipetting robot, together with the VOYAGER adjustable tip spacing

pipette, increases sample processing capacity, reduces the need for manual intervention

by laboratory personnel and fast tracks assay development for COVID-19 testing.

• Low retention GRIPTIPS and a low dead volume SureFlo reagent reservoir allow the loss

of costly reagents, such as one-step RT-PCR master mix, to be reduced.

• The simple and fast ASSIST PLUS pipetting robot combined with the easy-to-use

VIALAB software, offers immediate help for testing labs.

• While the current protocol uses 384 well plates, it can be readily adapted to 96 well format

to meet future needs.

• Thanks to the VIALAB software, the pipetting programs can be easily adapted to any

specific protocols and labware.

CHAPTER 3: Application Notes

For more information

and a list of materials

used, please refer to

our website.

57

3.3 Increase your sample screening and genotyping

assay throughput with the VOYAGER adjustable

tip spacing pipette

Discover the advantages of setting up a genotyping assay

or sample screening with the VOYAGER adjustable

tip spacing pipette

Laboratories are continually facing the challenge of

increasing throughput in the most efficient and economical

way, to meet the need to process more and more samples

per day. Traditionally, handling and manipulating samples

between different labware formats involves the use of single

channel pipettes, especially in screening applications and

genotyping assays, which is slow, inefficient and error prone.

INTEGRA’s VOYAGER adjustable tip spacing pipette has enabled

scientists from the Technical University of Munich (TUM) to benefit

from the enhanced productivity of a multichannel pipette, reducing

tedious liquid handling tasks.

Compared to fully automated solutions, it provides seamless liquid

transfers between different standardized and non-standardized microplates,

tube and gel chamber formats, and can be used without any special training.

Tip spacing can be simply changed one-handedly with the push of a button,

eliminating the need for any manual adjustments.

The various operating modes of the VOYAGER adjustable tip spacing pipette help to speed

up monotonous pipetting steps, eliminate sample transfer errors between different labware

formats, and reduce the risk of developing repetitive strain injuries.

CHAPTER 3: Application Notes

58 CHAPTER 3: Application Notes

Key benefits

• The VOYAGER’s motorized adjustable tip

spacing enables the user to benefit from

the enhanced productivity of an electronic

multichannel pipette throughout the entire

genotyping assay, processing samples

faster than with traditional single channel

pipettes and helping to eliminate sample

transfer errors between different labware

formats.

• Tip spacing can be adjusted on the fly with

the push of a button to match different

types of labware, allowing the easy transfer

of multiple reaction mix samples from

microcentrifuge tubes directly to 96 or

384 well plates, and gel pockets.

• The availability of a range of pipetting

modes makes the VOYAGER a very

versatile and affordable tool to speed up

and standardize pipetting protocols.

• New users quickly get accustomed to the

electronic pipette thanks to its intuitive

design and easy-to-use pipetting modes.

Experimental set-up

In this protocol, two VOYAGER 8 channel adjustable tip spacing pipettes are used for a

genotyping set-up. The genotyping assay is based on a PCR method with a subsequent gel

electrophoresis.

The following protocol consists of sample transfers from 1.5 ml microcentrifuge tubes into a

96 well plate, and from a 96 well PCR plate into an agarose gel for electrophoresis.

Overview of the steps:

1. Template transfer

2. Sample transfer into the agarose gel

CHAPTER 3: Application Notes 59

Figure 1: Adjust the tip spacing by aligning it against the empty 96 well plate and tube rack.

Step-by-step procedure

1. Template transfer

Transfer the templates into a 96 well plate.

Use a VOYAGER 8 channel 300 μl electronic pipette

with 300 μl sterile, filter GRIPTIPS. Select ‘Tip spacing’

in the main menu of the pipette to set the required

spacing. Choose ‘Positions: 2’ in the tip spacing menu

and set the tip spacing according to the 96 well plate

and the microcentrifuge tubes in the rack (Figure 1).

Once saved, the tip spacing is available at any time, for

any other pipetting modes.

After saving the tip spacing, select ‘Pipet’ mode in the

main menu. Set your required sample transfer volume

and pipette the templates from the 1.5 ml microcentrifuge tubes into the 96 well plate (Figure

2). By pressing left and right on the Touch Wheel interface, the tip spacing can be adjusted on

the fly to fit each labware format.

Tips:

• Use the Repeat Dispense mode to dispense several samples successively if duplicate or

triplicate samples are required.

• Use the Pipet/Mix mode if samples require mixing in the target wells. Settings like mixing

cycles, pipetting speeds and volumes can quickly be adjusted.

Figure 2: Sample transfer from a microcentrifuge tube rack

to a 96 well plate.

60

2. Sample transfer into the agarose gel

Transfer the PCR product into the agarose gel.

After PCR, use the VOYAGER 8 channel 125 μl

electronic pipette with 125 μl sterile, filter GRIPTIPS to

transfer the samples from the 96 well PCR plate into the

agarose gel for subsequent gel electrophoresis (Figure

3). As in step 1, choose ‘Positions: 2’ in the tip spacing

menu and set the tip spacing according to the 96 well

PCR plate and the agarose gel.

Set the required sample volume as described in step 1

and transfer the samples from the PCR plate into the

agarose gel.

Tips:

• A low dispensing speed (e.g. 4) helps uniform filling of the wells in the agarose gel.

• If you want a controlled blowin – rather than automatic – keep the run button pressed while

dispensing. Blowin will occur when the run button is released.

CHAPTER 3: Application Notes

Figure 3: PCR product transfer into the agarose gel.

Conclusion

• The VOYAGER adjustable tip spacing pipette has enabled TUM researchers using

different labware formats to benefit greatly from the enhanced productivity of a

multichannel pipette, processing assays much faster than using a single channel pipette.

The tip spacing can be changed onehandedly at the touch of a button to fit different

labware formats, such as PCR plates, tubes and gel pockets.

• Thanks to the intuitive interface, users quickly become accustomed to the electronic

pipette. The different pipetting modes make the VOYAGER adjustable tip spacing pipette

a versatile yet affordable tool for working with labware of varying sizes and formats.

• The VOYAGER adjustable tip spacing pipette increases the speed of sample testing

set-ups, and helps eliminate sample transfer errors between different labware formats

and reduce the risk of developing repetitive strain injuries.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 3: Application Notes 61

3.4 PCR product purification with QIAquick® 96

PCR Purification Kit and the VIAFLO 96

handheld electronic pipette

Semi-automated PCR product purification on the

VIAFLO 96 handheld electronic pipette

QIAquick 96 PCR Purification Kit is suitable for purifying

up to 10 μg of material for downstream applications,

such as sequencing, cloning, labeling and microarrays.

The kit facilitates the removal of impurities like primers,

unincorporated nucleotides, buffers, salts, mineral oils,

agarose and enzymes. The vacuum-driven process is

much faster than centrifugation, and gives high,

reproducible yields. It is important to avoid

cross-contamination in nucleic acid purification,

and QIAGEN's column design is optimized to limit

carryover of contaminants. Although QIAquick 96

provides a high throughput solution, the elution,

washing and binding steps are very laborious and

time consuming if performed manually. With

VIAFLO 96 handheld electronic pipette, the hands-on time

is reduced, as samples and reagents can be transferred to

all 96 wells at once. This enables rapid and efficient, high throughput PCR clean-up.

Key benefits

• VIAFLO 96 and VIAFLO 384 allow

simultaneous pipetting of up to 96 or 384

wells, respectively, maximize throughput

of PCR purification by allowing transfer

samples and reagents in a single step.

• The z-heights can be predefined, choosing

the optimal value to prevent accidental

scratching of the well membrane for more

consistent results.

• Custom programming of the PCR product

clean-up steps allows pipetting parameters,

such as aspiration or dispensing speeds, to

be predefined. Prompt messages guide the

user through the entire pipetting protocol,

which is especially useful when several

pre-wetting steps are included.

• The VIAFLO 96 or VIAFLO 384's handsfree

automatic mode ensures that the

PCR clean up protocols are performed

in the same way each time, maximizing

reproducibility.

62 CHAPTER 3: Application Notes

Overview: How to purify PCR products with

VIAFLO 96

Experimental set-up

This protocol describes how PCR products

are purified using a VIAFLO 96 handheld

electronic pipette with a two position stage and

the QIAGEN QIAquick® 96 PCR Purification

Kit. The following procedure is based on the kit

manufacturer's protocol for purification of 96

samples (up to 10 μg PCR products).

A 96 channel pipetting head (50-1250 μl) is

used together with 1250 μl short, low retention,

sterile, filter GRIPTIPS. Customized VIALINK

programs are provided to perform the binding,

washing and elution steps. Before starting,

ethanol (96-100 %) should be added to the

Buffer PE concentrate.

Overview of the purification steps:

1. Step 1: Binding

2. Step 2: Washing

3. Step 3: Elution

The initial set-up of the QIAvac 96 Vacuum Manifold consists of a waste tray on top of a QIAvac

base, followed by a QIAquick 96 well plate (pink) mounted on a QIAvac 96 top plate, as shown in

Figure 1.

The QIAvac has to be attached to a vacuum source (house vacuum or vacuum pump) that

generates negative pressure between 100 and 600 mbar.

Figure 1: Initial set-up of the vacuum manifold.

CHAPTER 3: Application Notes 63

Step-by-step procedure

1. Binding

Binding the DNA to the silica-gel membrane.

Load the 1250 μl short, low retention, sterile, filter GRIPTIPS

on the VIAFLO 96. Place a 150 ml automation friendly reagent

reservoir in position A. The QIAvac 96 Vacuum Manifold

should be placed on position B of the VIAFLO 96 in landscape

orientation. No plateholder is needed on position B where the

manifold is placed.

Important: The vacuum manifold should be aligned before

each run (Figure 2).

Begin by launching the custom VIALINK program 'Qiaquick_

purification_M'. The pipette will prompt the user to place Buffer

PM on position A, then air is aspirated. This ensures that every

single drop of the liquid can be dispensed later. The

VIAFLO 96 will then guide the user through the two pre-wetting

steps, starting with aspiration and dispensing 200 μl of Buffer

PM. After a second aspiration, the pipette will display the prompt

'Move the head out of buffer', before dispensing the final 200 μl of

Buffer PM. This is followed by a 20 second wait, giving the buffer

residues time to flow down to the tip and be dispensed.

After pre-wetting, the pipette aspirates 75 μl Buffer

PM (three times the volume of the PCR product).

The instrument then tells the user to remove the

reservoir from position A, and replace it with the

96 well plate containing the 25 μl of PCR products.

After dispensing, and four mixing steps, the

resulting mixture is transferred to the QIAquick plate

wells in two steps. It is then time to switch on the

vacuum source, as indicated by the pipette.

Tips:

• Pre-wetting the tips prior to pipetting prevents

droplets and dripping when pipetting volatile

liquids, such as isopropanol, which is one of the

constituents in Buffer PM.

• Low retention GRIPTIPS (Figure 3) are used for these pipetting steps to avoid dripping.

Figure 2: Alignment of the QIAvac 96 Vacuum

Manifold.

Figure 3: Low retention versus standard tips.

64 CHAPTER 3: Application Notes

2. Washing

Two-step purification of the PCR product.

Eject the used tips and load new 1250 μl short, low retention, sterile, filter GRIPTIPS on the

VIAFLO 96. Place a new 300 ml automation friendly reagent reservoir in position A. The

VIAFLO 96 will then prompt the user to pour Buffer PE into the reservoir, followed by a prewetting

step, which is necessary since the buffer contains ethanol. After pre-wetting, the pipette

will aspirate 900 μl of Buffer PE, and dispense it into QIAquick plate wells. The instrument will

then notify the user that is it time to turn on the vacuum pump. With the pump turned on, another

dose of the buffer is dispensed into the wells, followed by a 10 minute wait to dry the membrane

and remove all residual ethanol.

Important: The final drying step is crucial to remove residual ethanol prior to elution.

Residual ethanol in the elution buffer could inhibit downstream applications (e.g. PCR).

Tip: After this step, the manufacturer suggests tapping the plate on a stack of absorbent paper

to ensure that all residual buffer is removed.

3. Elution

Elution of DNA from the silica-gel membrane.

When prompted, start by replacing the waste tray with the

blue collection microtube rack provided, which contains

1.2 ml vessels (Figure 4a). Load new 1250 μl short, low

retention, sterile, filter GRIPTIPS, and place a new

150 ml automation friendly reagent reservoir in position A.

The instrument will then prompt the user to place Buffer

EB into the reservoir, aspirate 80 μl, and dispense it into

the QIAquick plate wells. After a 1 minute incubation, the

pipette tells the user to switch on the vacuum source for

5 minutes.

Tips:

• The purified PCR product could also be eluted in

a 96 well microplate. In this case, when replacing the

waste tray, the 96 well microplate has to be placed on

the empty blue collection tube rack (Figure 4b).

• For increased DNA concentration, decrease the elution

volume to 60 μl, as per QIAGEN's recommendations, in

the VIALINK software.

Figure 4: Elution into a) provided collection

microtubes or b) a 96 well microplate.

A)

B)

CHAPTER 3: Application Notes 65

Remarks

Vacuum manifold:

Alignment of the vacuum manifold is very important in this process. Adding marks on the deck

helps to reposition the manifold whenever needed. To check the position of the well plate on top

of the vacuum manifold, attach the tips manually to the pipette. The pipette tips should be in the

middle of the wells. If not, adjust the position of the vacuum manifold on the deck.

Automatic mode:

The VIAFLO 96 can also operate in hands-free automatic mode, allowing the user to have

more walk-away time and less interaction, which is highly beneficial when using the instrument

in a laminar flow cabinet. The customized automatic VIALINK program can be found on the

INTEGRA website.

Conclusion

• The VIAFLO 96 electronic handheld pipette allows fast and simple liquid transfers for high

throughput PCR product purification.

• Optimized pipette settings enable accurate sample and reagent transfer, without the tip

touching and scratching the QIAquick membrane.

• The VIAFLO 96 electronic handheld pipette's compact design takes up minimal space

and fits on any lab bench.

• The unique operating concept makes the VIAFLO 96 and VIAFLO 384 as easy to use as

a conventional electronic pipette.

• The QIAvac 96 manifold is easily placed on the instrument and allows the processing of

other kits using 96 well silica-membrane or filter plates.

• Another option for this application is the MINI 96, which is the most affordable 96 channel

option on the market.

For more information

and a list of materials

used, please refer to

our website.

66 CHAPTER 3: Application Notes

3.5 PCR purification with Beckman Coulter

AMPure XP magnetic beads and the VIAFLO 96

Automatic magnetic bead purification with the VIAFLO 96

handheld electronic pipette

Agencourt AMPure XP magnetic beads (Beckman Coulter) are an efficient

way to clean up samples for PCR, NGS, cloning and microarrays. The kit

provides a solution for medium to high throughput requirements when carried

out in a 96 well plate, but the protocol involves many washing and transfer

steps that make it tedious to perform manually. With the VIAFLO 96,

a handheld 96 channel electronic pipette, multistep protocols such as

PCR clean-up and DNA purification can be

performed quickly and efficiently, increasing

throughput tremendously by transferring

samples and reagents to all 96 wells at once.

Thanks to its unique operating concept,

the VIAFLO 96 remains as easy to use as

a traditional handheld pipette and can even

provide critical information (user-defined

prompts) about the protocol steps.

Key benefits

• The VIAFLO 96 enables transfer of

samples, reagents and wash solutions to

96 wells at once, increasing the throughput

of magnetic bead-based DNA purification

methods.

• The partial tip loading of the VIAFLO 96

allows purification of fewer than 96 DNA

samples if necessary; 8, 16, 24, 32, 40

or 48 GRIPTIPS can be loaded for easy

purification of different numbers of samples.

• The optimal immersion depth for removing

supernatant or adding liquid right onto

the samples is guaranteed by defining the

z-height of the VIAFLO 96.

• The Tip Align setting of the VIAFLO 96

automatically positions the tips in the center

of the wells of a 96 well plate, avoiding any

disturbance of the beads.

CHAPTER 3: Application Notes 67

Overview: How to automate PCR purification steps

with VIAFLO 96

The VIAFLO 96 handheld electronic pipette with a three position stage is used to purify DNA

with AMPure XP beads from Beckman Coulter. The following protocol is an example of a set-up

for 96 samples, where each well of a 96 well plate is filled with 10 μl of DNA sample and 18 μl

of AMPure XP beads, then further processed with the VIAFLO 96. The PCR purification can

be performed manually or semi-automated using the VIAFLO 96 in automatic mode.

Custom-made VIALINK programs are provided. The VIALINK programs are set up according

to the manufacturer’s protocol (AMPure XP Beckman Coulter).

Step-by-step procedure

1. Dispense AMPure XP beads into PCR tubes

Transfer AMPure XP beads from the stock solution into 12 PCR tubes placed in

a cooling block from INTEGRA.

Note: The cooling block is just used as a support in this instance, not for cooling down the

samples.

To ensure a homogenous stock solution, beads are thoroughly mixed by shaking/inverting until

the solution appears consistent in color. The beads are transferred into 12 PCR tubes using

the Repeat Dispense mode of a VIAFLO single channel 1250 μl electronic pipette. A customized

VIALINK program (AMP_Transfer1) is available to aid bead transfer.

For optimal pipetting, ensure beads are thoroughly mixed before each transfer. Mixing steps

can be defined by the number of cycles and the pipetting speed. Both influence the efficiency

of mixing and thus the quality of the

clean-up. Saving these parameters in the

pipetting program ensures that mixing is

always carried out as defined, yielding

consistent results. Insert a pre- and

post-dispense step to enhance accuracy

and precision while pipetting precious

reagents, such as AMPure XP beads.

Tip: The use of sterile, filter, low retention

GRIPTIPS ensures that every dispense

is as accurate as possible, with no loss of

beads or sample.

Figure 1: Transfer AMPure XP beads from the stock solution into 12 PCR

tubes.

68 CHAPTER 3: Application Notes

2. Transfer AMPure XP beads into the DNA samples

Transfer AMPure XP beads from the PCR tubes into a 96 well plate preloaded

with DNA samples.

Pipette the beads from the PCR tubes

into the 96 well plate using a VIAFLO

12 channel 50 μl electronic pipette. For

optimal pipetting, make sure the tips are

exchanged, and mix the beads thoroughly

before each transfer. A customized

VIALINK program (AMP_Transfer2) is

provided for this step.

Tip: Use low retention GRIPTIPS to

minimize loss of beads adhering to the

tip wall.

3. Mixing and binding of the AMPure XP beads

Mixing and binding of the magnetic beads to the PCR samples.

Load GRIPTIPS (position A) then select

and run the AMPure_XP_M program on

the VIAFLO 96. The samples are now

mixed 10 times by pipetting up and down

on position B. A five minute wait time

follows, timed by the VIAFLO 96, to allow

the DNA to bind to the beads.

Tip: Use the z-height setting of the

VIAFLO 96 to define the optimal tip

immersion depth. This prevents air

entering the tip during mixing and avoids

the pipette tip touching the bottom of the

plate. Setting the Tip Align support strength to 3 for positions A and B makes it more comfortable

to use the VIAFLO 96. These settings can be incorporated into the program so that they are not

forgotten.

Figure 2: Transfer AMPure XP beads from the PCR tubes into a 96 well

plate preloaded with DNA samples.

Figure 3: Mixing and binding of the magnetic beads to the PCR samples.

CHAPTER 3: Application Notes 69

4. Magnetic separation of the AMPure XP beads

Separating the magnetic beads from the PCR samples.

Note: Make sure new GRIPTIPS are loaded

before continuing the protocol to ensure

removal of the supernatant without bead

carryover.

A prompt on the pipette screen reminds the

user to move the sample plate from position

AB onto the 96 well magnet (position B) and

place an automation friendly reagent reservoir

for waste collection on position AB. After a two

minute incubation time, the beads form a ringshaped

structure and the solution becomes

clear. Load new GRIPTIPS before continuing the procedure to ensure accurate removal of

the supernatant without bead carryover. Follow the instructions on the pipette and aspirate

the supernatant slowly from the sample, dispensing it into the waste reagent reservoir

(position AB).

Tip: To avoid disturbing the ring of beads, the supernatant is aspirated slowly at speed 1.

Leave 5 μl of supernatant in the plate to prevent beads being drawn out during aspiration.

The z-height limit is again used to ensure that the beads are not disturbed during pipetting.

5. AMPure XP bead clean-up

Wash the magnetic beads twice with 70 % ethanol.

Place an automation friendly reagent reservoir

containing 70 % ethanol on position A and

change the GRIPTIPS before continuing

with the wash step. Follow the prompts on

the pipette. Pre-wet the GRIPTIPS with 70 %

ethanol. Then wash the samples with 70 %

ethanol. Repeat the washing step again as

indicated by the pipette.

Tip: Pre-wetting the GRIPTIPS with 70 %

ethanol ensures equilibration of the humidity

and the temperature between the air in the

pipette/tips and the sample/liquid. In-house testing has shown that low retention GRIPTIPS

prevent ethanol from dripping while traveling from one pipetting position to another.

Figure 4: Separating the magnetic beads from the PCR samples.

Figure 5: Wash the magnetic beads twice with 70 % ethanol.

70

6. Elute samples from the magnetic beads

Elute the purified samples from the magnetic beads by adding the elution

buffer.

As indicated by the pipette, replace the 70 %

ethanol reagent reservoir on position A with

an elution buffer reagent reservoir and move

the sample plate from the magnet (position B)

to position AB. Load new GRIPTIPS before

continuing with the protocol. After transferring

and thoroughly mixing the elution buffer with

the beads, the pipette prompts the user to

place the sample plate back onto the magnet

(position B). During the one minute incubation

time, place a new 96 well plate on position AB.

7. Transfer the sample eluates

Transfer the sample eluates into the new 96 well plate.

Note: Load new GRIPTIPS to ensure a clean

eluate transfer without bead carryover.

Continue with the same program, slowly and

carefully transferring the eluates from position

B into the new plate (position AB).

Tip: Optimizing pipette settings (aspiration

speed, volume and height) allows the volume

of the transferred eluate to be maximized

without carryover of beads. These settings

can be easily tweaked at any time. Performing

a test run with water before implementing any

new assay is an ideal way to optimize pipette settings.

Figure 6: Elute samples from the magnetic beads.

Figure 7: Transfer the sample eluates into the new 96 well plate.

CHAPTER 3: Application Notes

CHAPTER 3: Application Notes 71

Remarks

Automatic mode:

The VIAFLO 96 can also operate on its own,

enabling less user interaction, which in turn

improves ergonomics and reproducibility. This

also makes it even more ideal for use in tight

spaces, such as under a laminar flow cabinet.

Partial tip load:

If you are not working with a full set of 96

samples, the VIAFLO 96 is able to work with

any number of tips loaded, allowing purification

of smaller numbers of samples. Figure 8: Automatic mode and partial tip load.

Conclusion

• The VIAFLO 96 is perfectly suited to magnetic bead purification in a 96 well format. An

entire plate with 96 samples can be purified in a fraction of the time it would take with a

traditional pipette.

• Optimized tip immersion and pipette settings in combination with the use of low retention

GRIPTIPS allow maximum sample recovery at the end of the purification protocol.

• The VIAFLO 96 can guide the user through the entire protocol step by step, ensuring the

correct workflow and enhancing the reproducibility of results.

• The optional automatic mode of the VIAFLO 96 enables the instrument to operate on its

own to minimize pipetting errors, making it even more ideal for use under a laminar flow

cabinet.

For more information

and a list of materials

used, please refer to

our website.

72 CHAPTER 3: Application Notes

3.6 PCR purification with Beckman Coulter

AMPure XP magnetic beads and

the ASSIST PLUS

Automatic magnetic bead purification with

ASSIST PLUS pipetting robot

Agencourt AMPure XP beads (Beckman Coulter) are used

for DNA purification in a variety of applications, including PCR,

NGS, cloning and microarrays. The ASSIST PLUS pipetting

robot provides a solution for optimal bead

separation and maximized recovery of

precious samples. User guidance

throughout the entire protocol

ensures an error-free pipetting

procedure. Careful and accurate

handling of the magnetic beads

by the ASSIST PLUS leads to

superior reproducibility and consistency

during the experiment. Taken together, the

ASSIST PLUS provides researchers with an easy

and highly efficient way to purify DNA from PCR reactions using AMPure XP magnetic beads.

Key benefits

• The VIAFLO and VOYAGER electronic

pipettes, in combination with

ASSIST PLUS, provide unmatched

pipetting ergonomics.

• Optimal pipette settings, including tip

immersion depth, pipetting speeds and

angles, maximize reproducibility and

sample recovery.

• Exact positioning of the pipette tips in the

sample wells avoids the risk of disturbing

the ring of magnetic beads or bead

carryover.

• The ASSIST PLUS automates many steps

of a magnetic bead purification protocol

and guides the user through the remaining

manual operations to ensure an error-free

process.

CHAPTER 3: Application Notes 73

Overview: How to automate PCR purification steps

with ASSIST PLUS

The ASSIST PLUS is used to purify DNA samples using AMPure XP beads (Beckman Coulter).

The pipetting robot runs a VOYAGER 8 channel 125 μl electronic pipette with 125 μl sterile,

filter, low retention GRIPTIPS. The use of low retention GRIPTIPS guarantees optimal liquid

handling of viscous (AMPure XP buffer) and volatile (70 % ethanol) solutions.

Below is an example set-up for 24 samples, preparing 10 μl DNA samples (position B) with

18 μl of AMPure XP beads (position A). The pipetting programs were prepared according to the

manufacturer’s protocol (AMPure XP, Beckman Coulter) using VIALAB software.

The protocol is divided into two programs that guide the user through every step of the PCR

purification process.

• Program 1: Binding (AMP_BINDING)

• Program 2: Washing and elution (AMP_WASH_ELUTE)

Experimental set-up: Program 1

Deck position A: PCR 8 tube strip containing the AMPure XP

beads (Figure 1, blue), placed onto a cooling

block from INTEGRA. Note: the cooling block

is just used as a support in this instance, and

not for cooling down the samples.

Deck position B: 96 well plate with 24 DNA samples for

purification (Figure 1, green).

Deck position C: 96 well ring magnet.

74 CHAPTER 3: Application Notes

Figure 1: Pipetting schema, set-up for program 1.

A B C

Run program 1: transfer & binding

Select and run the AMP_BINDING program on the VOYAGER electronic pipette. The

ASSIST PLUS pipetting robot immediately starts the protocol.

1. AMPure XP transfer

Transferring AMPure XP beads from an 8 tube PCR strip to a 96 well plate

containing the DNA samples.

To ensure the AMPure XP buffer is homogenous, the beads are resuspended by pipetting up

and down 10 times before being transferred to the samples. The beads and DNA fragments

are thoroughly mixed together before the pipette automatically starts the timer for a 5 minute

incubation, ensuring optimal conditions for the DNA strands to bind onto the magnetic beads.

Tip: Using low retention GRIPTIPS rather than regular GRIPTIPS prevents the loss of AMPure

XP beads during the pipetting steps (see Figure 2).

VOYAGER 8 channel

125 μl

50/125 μl sterile, filter,

low retention GRIPTIPS

PCR 8-Tube Strip on cooling

plate – 200 μl

96 well plate Sapphire

– 200 μl

96 well plate Sapphire on 96 well ring

magnet – 200 μl

CHAPTER 3: Application Notes 75

Figure 2: The image highlights the advantages of using low retention GRIPTIPS versus regular

GRIPTIPS when pipetting AMPure XP beads.

Figure 3: The beads and DNA fragments are thoroughly mixed together before the incubation.

76 CHAPTER 3: Application Notes

2. Magnetic separation of the AMPure XP beads

Separating the magnetic beads from the PCR samples.

A message instructs the user to move the plate (position B) onto the magnet (position C).

Continue the program to start the timer. After a two minute incubation on the magnet the

beads form a ring in the sample well and the solution becomes clear. The program resumes

automatically, and the supernatant is removed. On completion of this step, the pipette prompts

the user to continue with the AMP_WASH_ELUTE program and to replace the labware on

position A with the 8 row polypropylene (PP) reagent reservoir containing the ethanol and

elution buffer.

Tip: The supernatant is aspirated slowly using the Tip Travel feature of the ASSIST PLUS

to avoid disturbing the ring of beads. The Tip Travel feature keeps the tip immersion depth

constant during aspiration and dispensing. 5 μl of supernatant remain in the plate to prevent

beads being drawn out during aspiration.

Figure 4: The ASSIST PLUS settings allow removal of the supernatant without any bead carryover.

CHAPTER 3: Application Notes 77

Experimental set-up: Program 2

Deck position A: The 96 well PCR cooling block is replaced by

an 8 row polypropylene (PP) reagent reservoir

filled with 70 % ethanol in row 1 (blue) and

elution buffer in row 2 (orange). Row 8 is used

for waste (purple).

Deck position B: Emtpy 96 well plate.

Deck position C: 96 well ring magnet and 96 well plate with 24

DNA samples for purification (green).

Figure 5: Pipetting schema, set-up for program 2.

VOYAGER 8 channel

125 μl

50/125 μl sterile, filter,

low retention GRIPTIPS 8 row reagent reservoir 96 well plate Sapphire

– 200 μl

96 well plate Sapphire on 96 well ring

magnet – 200 μl

A B C

78 CHAPTER 3: Application Notes

Run program 2: Washing & elution

Start the AMP_WASH_ELUTE program on the VOYAGER electronic pipette. The

ASSIST PLUS washes the beads twice by automatically adding and removing ethanol.

3. Magnetic bead clean-up

Washing the magnetic beads twice with 70 % ethanol.

The programmed pipette settings allow the beads to be washed without disturbing the bead

ring. At the end of the second washing step, all the ethanol is removed. If necessary, an

additional drying time can easily be added using VIALAB software.

Tip: The use of low retention GRIPTIPS prevents ethanol from dripping while traveling from

position A to position C (see Figure 6).

Figure 6: The image highlights the advantages of using low retention GRIPTIPS (left) versus regular

GRIPTIPS (right) when pipetting ethanol.

4. Elute samples from the magnetic beads

Eluting the samples from the magnetic beads by adding an elution buffer.

The pipette prompts the user to move the reaction plate from the magnet (position C) to position

B. Continuing the protocol, the ASSIST PLUS transfers the elution buffer to the DNA samples

bound to the magnetic beads (position B, orange). After mixing carefully and thoroughly 10

times, the pipette prompts the user to place the 96 well plate on the magnet (position C).

CHAPTER 3: Application Notes 79

5. Transfer the sample eluates

Transferring the sample eluates into a new 96 well plate.

As indicated by the pipette, place a new 96 well plate onto position B and continue the program.

The sample eluates are then transferred into the new plate automatically.

Tip: Optimized pipette settings (aspiration speed, volume, height, tip travel and tip touch) allow

the volume of eluate transferred to be maximized without carryover of beads (see Figure 6).

A tip touch after the transfer removes droplets that may still cling to the end of the pipette tips.

Pipetting heights on the ASSIST PLUS can be fine-tuned at any time. Performing a test run with

water before implementing any new assay is an ideal way to optimize pipette settings.

Results

Figure 7: Magnetic beads are clearly visible in the 96 well plate with no supernatant remaining.

80 CHAPTER 3: Application Notes

Figure 8: No carryover of beads is observed in the eluate.

Conclusion

• Magnetic bead purifications can be easily automated on the ASSIST PLUS pipetting

robot.

• Optimized tip immersion and pipette settings together with the use of low retention

GRIPTIPS allow maximum sample recovery at the end of the purification protocol.

• The pipette loaded onto the ASSIST PLUS prompts the user when needed, eliminating

the risk of human errors.

• VIALAB programs can be easily adapted to specific labware.

• Prolonged pipetting tasks lead to repetitive strain injury. This can be avoided by

automating these steps with the ASSIST PLUS.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 4: Customer Testimonials 81

CHAPTER 4:

Customer testimonials

Our range of innovative liquid handling products has helped countless laboratories to achieve

PCR success, improve their throughput and further their ground-breaking research. But don’t

just take our word for it! Here are a few stories from our satisfied customers, demonstrating why

INTEGRA Biosciences is the right choice for PCR pipetting solutions and labware.

4.1 INTEGRA pipettes – the obvious choice for

start-up PCR labs

The gradual reopening of the world following the pandemic has led to an unprecedented

demand for COVID-19 testing, with schools, universities and workplaces relying on negative

PCR tests to continue operating. Matrix Diagnostics – a dedicated COVID-19 testing lab in

California – is helping to fulfill this critical need, relying on INTEGRA’s EVOLVE and MINI 96

pipettes to streamline and accelerate PCR workflows.

PCR-based diagnostic testing is a well-established technique in clinical labs around the world,

and this method has been brought to the attention of every household as the gold standard for

COVID-19 testing. However, the public is less aware that the sensitivity of this technique makes

it time-consuming and troublesome to perform without the right tools, as it is very sensitive to

pipetting errors and cross-contamination.

Founded in January 2021, Matrix Diagnostics

was established to meet the growing demand

for PCR testing in the San Francisco Bay Area,

and the newly formed team understood the need

for effective pipetting solutions from the outset.

Fady Ettnas, Lab Manager at Matrix Diagnostics,

explained: “We realized that, to meet the

anticipated demand for testing, we would have to

turnover between 2000 and 5000 samples every

day. This seemed like an impossible task for a new

lab with limited resources but, after implementing

INTEGRA’s pipettes in our lab, we quickly

alleviated the pipetting bottlenecks, putting us on

track to achieve our targets.”

Photo courtesy of Matrix Diagnostics

82

Evolving workflows

“Our protocols involve a range of repetitive

pipetting steps – including mixing reagents

and serial dilutions – for thousands of

samples a day, which has the potential

to be a cumbersome and error-prone

task,” Fady continued. “We therefore

chose INTEGRA’s EVOLVE manual

pipettes and MINI 96 portable electronic

pipettes to improve the reproducibility

and productivity of our workflows. We

have a number of single channel EVOLVE

pipettes, covering volumes ranging from

0.2 to 5000 μl, as well as 8, 12, and 16

channel models. What I like most about

EVOLVE is its ergonomic design and

ability to set volumes in a flash. The

unique design of INTEGRA’s GRIPTIPS also means that they never leak or fall off, avoiding

cross-contamination and maintaining sterility. We also use the compact MINI 96 extensively,

which is especially well suited to PCR set-up. It saves a lot of time and effort – around 15

minutes per cycle – when performing the wash steps. And because we run more than 25 cycles

every day, this is a huge saving, allowing us to process a much higher number of samples. It is

a perfect and affordable solution for our needs.”

A long-term investment

The benefits of these pipettes to users, particularly in terms of preventing physical strain

caused by repeated pipetting actions, are a priceless advantage. “I think the pipettes are a

great investment with huge returns, allowing the team to process more samples and improving

their pipetting experience. The company’s customer service is quick, responsive and helpful

and, crucially, the team was able to advise us on the right choice of pipettes to meet our

workload and objectives.”

Planning future with INTEGRA

“Currently, we are only offering COVID-19 tests, but we plan to expand to include other tests

including sexually transmitted diseases, urinary tract infections and flu, and we know that we

will need to automate our workflow. We will need something flexible and incredibly efficient and,

therefore, we are planning to acquire an ASSIST PLUS pipetting robot. I like all the INTEGRA

products that I’ve used, and have rarely encountered even minor technical issues. I think they

are the most obvious pipetting choice for both for start-ups and established lab set-ups, and are

well worth the investment,” Fady concluded.

Photo courtesy of Matrix Diagnostics

CHAPTER 4: Customer Testimonials

83

Photo courtesy of Harvard Medical School

4.2 A better qPCR pipetting experience

Manual pipetting can be a major bottleneck for research laboratories, especially when they face

the challenge of combining accurate results with high throughput. Like all repetitive tasks that

require precise actions, filling multiwell plates by hand is time consuming, and physically and

mentally draining, which can lead to errors. When Daisy Shu joined the Saint-Geniez laboratory

at Harvard Medical School, her experience was quite different, thanks to the INTEGRA VIAFLO

electronic pipettes.

From patients to pipettes

After graduating in optometry from the University of New South

Wales in Sydney, Daisy worked as an optometrist for two years

before deciding to pursue a PhD in cataract research at the

University of Sydney. She explained: “The move from my usual

clinical work with patients to research was a big change for me,

as I had to dive deep into molecular biology. I didn't even know

how to use a pipette back then! Cataracts – clouding of the

eye’s lens – are a leading cause of blindness worldwide, and I

studied their formation and ways to prevent that happening. My

focus was on transforming growth factor beta (TGF-β), which

has an important role in cancer metastasis, but is also relevant

for certain types of cataracts. I looked at the different signaling

pathways it activates and how those pathways interlink.”

Daisy completed her PhD in January 2019, and straight

afterwards flew to Boston to work as postdoctoral fellow in the

Saint-Geniez laboratory, continuing her research into eye health.

Here, she was able to apply her knowledge of TGF-β to agerelated

macular degeneration (AMD). Daisy continued: “I'm now

looking at how TGF-β causes the retinal mitochondria to change morphology and become

dysfunctional, altering cellular metabolism. The research is still at an early stage, so we're

mainly trying to understand how to prevent AMD, but the end goal is to find a cure.”

A better pipetting experience

At Harvard, Daisy was introduced to VIAFLO electronic pipettes, which were a complete

contrast to the large, fully automated pipetting workstation she had used during her PhD

research. The laboratory was already using two VIAFLO pipettes – a 125 μl eight channel

pipette and a 12.5 μl single channel version – and their flexibility compared to the automated

workstation dramatically improved her pipetting experience. “Complete automation on a large

workstation has its place, but there are downsides,” said Daisy. “You have to program every

CHAPTER 4: Customer Testimonials

84

single step perfectly before you can click one button and run the

protocol, and the process of fine-tuning takes a long time.”

“I found the VIAFLO pipettes amazing. A lot of our work is PCRbased,

performed in 384 well plates, and the VIAFLO pipettes

are real lifesavers. I use the 8 channel VIAFLO for most qPCR

liquid transfers, and the single channel pipette to add the

primers. Once you've made your master mixes and programmed

the pipette, it's really fast; it only takes me 20 minutes to

do a complete 384 well plate. When I was using the robotic

workstation in Sydney, I used to think that doing a qPCR was

really a big deal. Now, with the INTEGRA pipettes, it's just

so easy.”

VIAFLO pipettes provide a choice of pipetting modes and allow

easy adjustment of parameters such as volume and speed, as

well as providing pre-set programs and the option for custom

workflows. This helps laboratories to reduce errors and increase

throughput and reproducibility regardless of the users’ pipetting

experience. For Daisy, VIAFLO electronic pipettes have become the standard for how pipetting

should be: “In any pipetting workflow, you have to get every step right first time, otherwise you’d

end up having to troubleshoot the assay and do it again. I'm really surprised when I hear people

from other labs say they pipette each well individually with manual single channel pipettes. I’m

sure that would take forever compared to electronic pipetting, and my eyes would really suffer.

The VIAFLOs make everything easy. I love the color coding – it makes it so simple to match the

right tip to the right pipette – and the instrument can even be set to alert you when you need to

pipette again.”

CHAPTER 4: Customer Testimonials

Photo courtesy of Harvard Medical School

85

4.3 COVID-19 – Accelerate your PCR set-up

The emergence and outbreak of the novel coronavirus SARS-CoV-2 (COVID-19) has placed

unprecedented demands on laboratories testing patient samples for COVID-19, leaving

scientific staff to contend with a spiraling influx of COVID-19 samples and a rapid, continuous

growth in workload. Among the challenges faced by the Microbiology and Molecular Pathology

Department at Sullivan Nicolaides Pathology (SNP) – part of the Sonic Healthcare Group

– in Brisbane, Australia, is the increased pressure on laboratory automation used for both

coronavirus and pre-existing respiratory virus panel testing.

As a result of the coronavirus pandemic, SNP found itself analyzing extreme numbers of

samples, which exhausted the capacity of its automation platforms. At the same time, staff

were faced with a need to spend more time working up new virus testing protocols, which

were often performed manually or using semi-automated methods to fast track test response

times, leaving them prone to increased ergonomic strain. There was a clear need for additional

automated liquid handling instruments to increase sample processing capacity, reduce manual

intervention by laboratory analysts and fast track assay development for COVID-19 sample

testing.

Working together

In early March 2020, Kelly Magin and James Sundholm from

INTEGRA’s Australian distributor, BioTools Pty Ltd, partnered

with Shane Byrne, Scientific Department Head, Microbiology and

Molecular Pathology Department, SNP, to support COVID-19 testing

of patient samples using the ASSIST PLUS pipetting robot. An

ASSIST PLUS automated pipetting protocol was developed and

validated, enabling samples to be prepared in low volume, 384 well

plates for subsequent processing on a rapid, high throughput,

plate-based, real-time PCR amplification and detection instrument.

A VOYAGER adjustable tip spacing pipette and low retention

GRIPTIPS were used to transfer one-step RT-PCR master mix from a

low dead volume (<20 μl) SureFlo 10 ml reagent reservoir into a 384

well plate. The VOYAGER pipette also allowed automatic transfer

and reformatting of nucleic acid template extracted from combined

nasopharyngeal/oropharyngeal flocked swab(s) or sputum samples,

from 4 x FluidX™ 1.0 ml 96 format tube racks into the 384 well plate. The total PCR reaction

volume was reduced to 10 μl; 7.5 μl one-step RT-PCR master mix and 2.5 μl of nucleic acid

template. This miniaturization doubled the available testing capacity and simultaneously

reduced consumption of expensive one-step RT-PCR reagents of dwindling availability, with

associated cost savings.

Photo courtesy of Sullivan Nicolaides

Pathology

CHAPTER 4: Customer Testimonials

86

Defining success

SNP successfully validated the automated

protocol against its existing manual

processing method, performed using a

handheld electronic pipette. The results

were shown to be reproducible, precise

and accurate, with no contamination

observed in either the control or patient

samples. The compact, easy-to-use

ASSIST PLUS pipetting robot, complete

with validated protocol, was fully deployed

within five working days. While the current

protocol uses 384 well plates, it can be

readily adapted to 96 well format to meet

future needs.

4.4 Reducing protocol time for PCR using

96 channel pipette

Implementing an INTEGRA VIAFLO 96 electronic pipette has enabled the Virus- and Prion

Validation (VPV) Department at Octapharma Biopharmaceuticals GmbH, (Frankfurt, Germany)

to reduce the time taken to undertake PCR assays by greater than 60 %.

Since its foundation in 1983, Octapharma has been committed to patient care and medical

innovation. Its core business is the development and production of human proteins from human

plasma and human cell-lines.

The VPV Department has been set-up to investigate pathogen inactivation and removal steps

along the manufacturing processes. Among other techniques, multi-step 96 well format PCR

assays were developed, which involve three washing steps twice in the protocol. To undertake

their PCR assay more efficiently, Octapharma sought a system that enabled reproducible and

accurate liquid handling in the 96 well format and was able to completely remove residual liquid

as well as avoid well-to-well contamination.

Dr. Andreas Volk, a research scientist at Octapharma Biopharmaceuticals commented: "The

classical liquid handling solutions, fully automated robots or ELISA plate washers were either

too costly or prone to cross contamination in a PCR assay." He added: "When we tested the

INTEGRA VIAFLO 96 channel pipette, it fully met our requirements as it enabled mediumthroughput

liquid handling while minimizing cross-contamination. Additionally, the

Photo courtesy of Sullivan Nicolaides Pathology

CHAPTER 4: Customer Testimonials

87

VIAFLO 96 electronic pipette provided all the

adjustment options, which we had been used

to with manual pipettes, plus a specified tip

immersion depth for each pipetting step. With

our PCR protocol, which involves ten full liquid

transfers per plate, we now only use half the

amount of pipette tips as we can use the same

tips for liquid addition and aspiration in each

washing step. VPV Department staff has found

using the VIAFLO 96 benchtop pipette highly

intuitive and the overall time required for our

PCR washing procedures has been reduced to

approximately one third of the original time."

The INTEGRA VIAFLO 96 is a handheld 96

channel electronic pipette that has struck a

chord with scientists looking for fast, precise

and easy simultaneous transfer of

96 samples from microplates without the

cost of a fully automated system. The

VIAFLO 96 was designed to be handled just

like a standard handheld pipette – a fact

borne out by consistent end user feedback

that no special skills or training are required to

operate it. Users immediately benefit from the

increased productivity delivered by their VIAFLO 96. Fast replication or reformatting of 96 and

384 well plates and high precision transferring of reagents, compounds and solutions to or from

microplates with the VIAFLO 96 is as easy as pipetting with a standard electronic pipette into

a single tube. Four pipetting heads with pipetting volumes up to 12.5 μl, 125 μl, 300 μl or

1250 μl are available for the VIAFLO 96. These pipetting heads are interchangeable within

seconds enabling optimal matching of the available volume range to the application performed.

For 384 well pipetting, an enhanced version is available with VIAFLO 384. It features

384 channel pipetting heads in the volume range of 12.5 μl and 125 μl and is compatible with

96 channel pipetting heads.

Dr. Andreas Volk, Octapharma Biopharmaceuticals

CHAPTER 4: Customer Testimonials

88

CHAPTER 5:

Conclusion

So, there you have it, a full run down of PCR. By now, you should have all the information you

need to become a PCR pro, but if you’d still like to learn more about this interesting topic, we

have a wealth of articles on our website. Whatever your PCR requirements, we at INTEGRA

Biosciences are always available to answer your questions and provide you with the best

workflow solutions.

CHAPTER 5: Conclusion

89

CHAPTER 6:

References

1.1 The complete guide to PCR

1. Crow, E. (2012). Mind Your P's And Q's: A Short Primer On Proofreading Polymerases.

https://bitesizebio.com/8080/mind-your-ps-and-qs-a-short-primer-on-proofreadingpolymerases

2. Kim, S. W. et al. (2008). Crystal structure of Pfu, the high fidelity DNA polymerase from

Pyrococcus furiosus. International Journal of Biological Macromolecules, 42(4), 356-

361. https://doi.org/10.1016/j.ijbiomac.2008.01.010

3. ThermoFisher Scientific (n.d.). PCR Setup – Six Critical Components to Consider.

https://www.thermofisher.com/ch/en/home/life-science/cloning/cloning-learningcenter/

invitrogen-school-of-molecular-biology/pcr-education/pcr-reagents-enzymes/

pcr-component-considerations.html

4. AAT Bioquest (2020). What is the function of MgCl2 in PCR?

https://www.aatbio.com/resources/faq-frequently-asked-questions/What-is-thefunction-

of-MgCl2-in-PCR

5. Lorenz, T. C. (2012). Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting

and Optimization Strategies. Journal of Visualized Experiments, 63, e3998.

https://doi.org/10.3791/3998

6. Merck (n.d.). Polyermase Chain Reaction.

https://www.sigmaaldrich.com/CH/en/technical-documents/technical-article/

genomics/pcr/polymerase-chain-reaction

7. Viana, R. V., Wallis, C. L. (2011). Good Clinical Laboratory Practice (GCLP) for

Molecular Based Tests Used in Diagnostic Laboratories. In Akyar, I. (Ed.), Wide

Spectra of Quality Control (29-52). InTech.

https://cdn.intechopen.com/pdfs/23728/InTech-Good_clinical_laboratory_

practice_%20gclp_for_molecular_based_tests_used_in_diagnostic_laboratories.pdf

8. Ogene M. (2021). How does ddPCR work?

https://mogene.com/how-does-ddpcr-work

9. ThermoFisher Scientific (2016). Real-time PCR handbook.

https://www.ffclrp.usp.br/divulgacao/emu/real_time/manuais/Apostila%20qPCRHandbook.

pdf

CHAPTER 6: References

90

10. Prediger, E. (2017). Digital PCR (dPCR) – What is it and why use it?

https://eu.idtdna.com/pages/technology/qpcr-and-pcr/digital-pcr

11. Bio-Rad Laboratories (n.d.). Introduction to Digital PCR.

https://www.bio-rad.com/en-uk/life-science/learning-center/introduction-to-digital-pcr

12. Bio-Rad Laboratories (n.d.). Digital PCR and Real-Time PCR (qPCR) Choices for

Different Applications.

https://www.bio-rad.com/en-uk/life-science/learning-center/digital-pcr-and-real-timepcr-

qpcr-choices-for-different-applications

13. Schoenbrunner, N. J. et al. (2017). Covalent modification of primers improves PCR

amplification specificity and yield. Biology Methods and Protocols, 2(1).

https://doi.org/10.1016/j.ijbiomac.2008.01.010

14. Merck (n.d.). Hot Start PCR.

https://www.sigmaaldrich.com/CH/en/technical-documents/technical-article/

genomics/pcr/hot-start-pcr

15. Parichha, A. (2021). Nested PCR || Principle and usage.

https://www.youtube.com/watch?v=nHCjgo2Ze0o

16. New England Biolabs (n.d.). FAQ: What is touchdown PCR?

https://international.neb.com/faqs/0001/01/01/what-is-touchdown-pcr

17. Parichha, A. (2021). Touch down PCR.

https://www.youtube.com/watch?v=s9oV2-53esA

18. Cheriyedath, S. (2018). History of Polymerase Chain Reaction (PCR).

https://www.news-medical.net/life-sciences/History-of-Polymerase-Chain-Reaction-

(PCR).aspx

19. Arney, K. (2020). The Story of PCR.

https://geneticsunzipped.com/news/2020/11/3/the-story-of-pcr

20. Biosearch Technologies (2022). Taq facts.

https://blog.biosearchtech.com/thebiosearchtechblog/bid/48174/taq-facts

21. National Museum of American History (n.d.). Mr. Cycle, Thermal Cycler.

https://americanhistory.si.edu/collections/search/object/nmah_1000862

CHAPTER 6: References

91

1.2 Simple PCR tips that can make or break your success

1. Cheriyedath, S. (2018). History of Polymerase Chain Reaction (PCR).

https://www.news-medical.net/life-sciences/History-of-Polymerase-Chain-Reaction-

(PCR).aspx

2. Seeding Labs (2019). How To: PCR Calculations.

https://www.youtube.com/watch?v=CnQV5_CEvAo

3. McCauley, B. (2020). Setting Up PCR Reactions.

https://brianmccauley.net/bio-6b/6b-lab/polymerase-chain-reaction/pcr-setup

4. New England Biolabs (n.d.). Guidelines for PCR Optimization with Taq DNA

Polymerase.

https://international.neb.com/tools-and-resources/usage-guidelines/guidelines-forpcr-

optimization-with-taq-dna-polymerase

5. Lorenz, T. C. (2012). Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting

and Optimization Strategies. Journal of Visualized Experiments, 63, e3998.

https://doi.org/10.3791/3998

6. Gold Biotechnology (2020). How To: PCR Master Mixes.

https://www.youtube.com/watch?v=LSfvCJ9gUQU

1.3 Setting up a PCR lab from scratch

1. Bustin, S. A., Benes, V., Garson, J. A., et al (2009). The MIQE Guidelines: Minimum

Information for Publication of Quantitative Real-Time PCR Experiments. Clinical

Chemistry, 55(4), 611–622.

https://doi.org/10.1373/clinchem.2008.112797

2. National Human Genome Research Institute (2020). Polymerase Chain Reaction

(PCR) Fact Sheet.

https://www.genome.gov/about-genomics/fact-sheets/Polymerase-Chain-Reaction-

Fact-Sheet

3. Viana, R. V., Wallis, C. L. (2011). Good Clinical Laboratory Practice (GCLP) for

Molecular Based Tests Used in Diagnostic Laboratories.

https://cdn.intechopen.com/pdfs/23728/InTech-Good_clinical_laboratory_practice_

gclp_for_molecular_based_tests_used_in_diagnostic_laboratories.pdf

4. Redig, J. (2014). The Devil is in the Details: How to Setup a PCR Laboratory.

https://bitesizebio.com/19880/the-devil-is-in-the-details-how-to-setup-a-pcrlaboratory

5. Mifflin, T. E. (n. d.). Setting Up a PCR Laboratory.

https://pubmed.ncbi.nlm.nih.gov/21357132/

CHAPTER 6: References

92

6. Gu, M. (n. d.). Molecular Laboratory Design And Its Contamination Safeguards.

https://www.scimmit.com/molecular-laboratory-design-and-its-contaminationsafeguards

7. Lee, R. (2015). Molecular Laboratory Design, QA/QC Considerations.

https://www.aphl.org/programs/newborn_screening/Documents/2015_Molecular-

Workshop/Molecular-Laboratory-Design-QAQC-Considerations.pdf

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work

1. Bustin, S. A., Benes, V., Garson, J. A. et al. (2009). The MIQE guidelines: minimum

information for publication of quantitative real-time PCR experiments. Clinical

Chemistry, 55(4), 611-622.

https://doi.org/10.1373/clinchem.2008.112797

2. Rutledge, R. G., Côté, C. (2003). Mathematics of quantitative kinetic PCR and the

application of standard curves. Nucleic Acids Research, 31(16).

https://www.gene-quantification.de/rudledge-2003.pdf

3. Applied biological materials (2016). Polymerase chain reaction (PCR) – Quantitative

PCR (qPCR).

https://www.youtube.com/watch?v=YhXj5Yy4ksQ

4. Nagy, A., Vitásková, E., Černíková, L. et al. (2017). Evaluation of TaqMan qPCR

system integrating two identically labelled hydrolysis probes in single assay. Scientific

reports, 7.

https://doi.org/10.1038/srep41392

5. Bradburn, S. (n.d.). How to calculate PCR primer efficiencies.

https://toptipbio.com/calculate-primer-efficiencies

6. Bio-Rad (n.d.). qPCR assay design and optimization.

https://www.bio-rad.com/en-ch/applications-technologies/qpcr-assay-designoptimization?

ID=LUSO7RIVK

7. University of Western Australia (2016). Melt curve analysis in qPCR experiments.

https://www.youtube.com/watch?v=FvJnXKzejSQ

8. Bio-Rad (2011). Real time QPCR data analysis tutorial.

https://www.youtube.com/watch?v=GQOnX1-SUrI

9. Bio-Rad (2011). Real time QPCR data analysis tutorial (part 2).

https://www.youtube.com/watch?v=tgp4bbnj-ng

10. Kannan, S. (2021). 4 easy steps to analyze your qPCR data using double delta Ct

analysis.

https://bitesizebio.com/24894/4-easy-steps-to-analyze-your-qpcr-data-using-doubledelta-

ct-analysis

CHAPTER 6: References

93

1.5 How to design primers for PCR

1. Benchling (n.d.). Primer Design.

https://www.benchling.com/primers

2. Addgene (n.d.). How to Design a Primer.

https://www.addgene.org/protocols/primer-design

3. PREMIER Biosoft (n.d.). PCR Primer Design Guidelines.

http://www.premierbiosoft.com/tech_notes/PCR_Primer_Design.html

4. Merck (n.d.). Oligonucleotide Melting Temperature.

https://www.sigmaaldrich.com/CH/en/technical-documents/protocol/genomics/pcr/

oligos-melting-temp

5. Integrated DNA Technologies (n.d.). How do you calculate the annealing temperature

for PCR?

https://eu.idtdna.com/pages/support/faqs/how-do-you-calculate-the-annealingtemperature-

for-pcr

CHAPTER 6: References

INTEGRA Biosciences AG

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HOW TO BECOME A PCR PRO

The polymerase chain reaction (PCR) is a key life sciences technique. It has been used in

molecular biology – including molecular diagnostics – for many years, and a number of different

types, for example, RT-PCR, qPCR, vPCR and ddPCR have been developed over time.

Today, PCR is a vital tool for the detection of pathogens, such as the SARS-CoV-2 virus, and

is essential for genotyping and NGS library preparation. However, PCR is well known for being

difficult to run successfully and several parameters must be considered when planning the PCR

protocol.

We have therefore compiled this eBook – consisting of in-depth educational articles, relevant

app notes and customer testimonials – to help you understand how PCR works, and what needs

to be considered to perform effective PCR reactions. We also demonstrate how our solutions

can help you to enhance the throughput of your lab, and become a PCR pro in no time.

Dr Éva Mészáros

Application Specialist

eva.meszaros@integra-biosciences.com

Anina Werner

Content Manager

anina.werner@integra-biosciences.com

FOREWORD

TABLE OF CONTENTS

CHAPTER 1: What you need to know about PCR

1.1 The complete guide to PCR 2

1.2 Simple PCR tips that can make or break your success 15

1.3 Setting up a PCR lab from scratch 20

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work 24

1.5 How to design primers for PCR 32

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 38

CHAPTER 3: Application notes

3.1 Efficient and automated 384 well qPCR set-up with the ASSIST PLUS pipetting robot 42

3.2 Automated RT-PCR set-up for COVID-19 testing 49

3.3 Increase your sample screening and genotyping assay throughput with the VOYAGER 57

adjustable tip spacing pipette

3.4 PCR product purification with QIAquick® 96 PCR Purification Kit and the 61

VIAFLO 96 handheld electronic pipette

3.5 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 66

VIAFLO 96

3.6 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 72

ASSIST PLUS

CHAPTER 4: Customer testimonials

4.1 INTEGRA pipettes – the obvious choice for start-up PCR labs 81

4.2 A better qPCR pipetting experience 83

4.3 COVID-19 – Accelerate your PCR set-up 85

4.4 Reducing protocol time for PCR using 96 channel pipette 86

CHAPTER 5: Conclusion 88

CHAPTER 6: References 89

2

CHAPTER 1:

What you need to know about PCR

In this chapter, we will cover topics such as PCR’s fascinating history, its mechanism and

different variations, and techniques for troubleshooting common issues you may encounter.

We’ll also go through tips for establishing a PCR lab, as well as a comprehensive overview of all

things related to qPCR and primer design.

1.1 The complete guide to PCR

Polymerase chain reaction (PCR) methods have been carried out in labs around the world since

the 1980s, opening the door for an array of new applications, such as genetic engineering,

genotyping and sequencing. In this article, we take a deep dive into this fascinating technique

by explaining its mechanism, exploring its history, looking into the different types of PCR,

discussing troubleshooting tips and much more.

CHAPTER 1: What you need to know about PCR

3

What is PCR?

The polymerase chain reaction (PCR) is a fast and inexpensive technique for amplifying a DNA

sequence of interest. It consists of three steps:

• Denaturation: The sample is heated to separate the DNA into two single strands.

• Annealing: The temperature is lowered to allow primers to anneal to specific single-stranded

DNA segments, flanking the sequence to be amplified.

• Extension: The temperature is raised to the optimum working temperature of the polymerase

enzyme, which then makes a complementary copy of the DNA sequence of interest.

One such repetition or 'thermal cycle' theoretically doubles the amount of the DNA sequence of

interest present in the reaction. Typically, 25 to 40 cycles are performed – resulting in millions

or even billions of DNA copies – depending largely on the amount of DNA in the starting sample

and the number of amplicon copies needed for post-PCR applications.

The three steps of a PCR reaction are carried out automatically by a thermal cycler, but can

only be successful if the master mix has been correctly prepared. The following sections

explain the components that make up the master mix and how they interact with the template

DNA during thermal cycling.

PCR master mix components

The PCR master mix consists of six components:

• PCR-grade water: Certified to be free of contaminants, nucleases and inhibitors.

• dNTPs: Containing equal concentrations of the four nucleotides (dATP, dCTP, dGTP and

dTTP), which are the 'building blocks' to create complementary copies of the DNA sequence

of interest.

• Forward and reverse primers: Short, single-stranded DNA sequences that anneal

specifically to the plus and minus strands of the template DNA, flanking the sequence to

be amplified. For some assays – such as protocols amplifying much-studied genes or DNA

sequences of common bacteria – ready-to-use primers can be purchased. However, many

experiments require custom PCR primers tailored to the region of interest of the template DNA

and the reaction conditions.

CHAPTER 1: What you need to know about PCR

4

• DNA polymerase: Taq-polymerase is the most commonly used enzyme for PCR reactions.

It uses dNTPs to create complementary copies of the DNA sequence of interest. For some

applications, such as mutagenesis, Taq-polymerase is not accurate enough and the use

of high fidelity polymerases is recommended. Just like Taq-polymerase, they sometimes

add an incorrect nucleotide when replicating the template DNA but, as they have a 3' to

5' exonuclease activity, they 'proofread' the newly synthesized strands and correct any

mistakes.1 This proofreading step is highly beneficial for accuracy but it also slows down PCR

reactions, and high fidelity polymerases (also called slow polymerases) therefore need about

twice the time of Taq-polymerase to create a complementary DNA strand. The most popular

high fidelity DNA polymerase is Pfu-polymerase.2

• Buffer: Provides a suitable environment for the DNA polymerase, with a pH between 8.0

and 9.5.3

• Magnesium chloride: Increases the activity of the DNA polymerase and helps primers

to anneal to the template DNA for a higher amplification rate.4 This cofactor is sometimes

included in the buffer in a sufficient concentration.5

The template DNA , which may be genomic DNA (gDNA), complementary DNA (cDNA) or

plasmid DNA (pDNA), is then added after master mix preparation.

The 3 steps of PCR

After preparing the PCR master mix and adding the template DNA samples to it, you can load

your reaction tubes, PCR strips or microplates into the thermal cycler. They will then go through

the following steps:

• Denaturation: The thermal cycler first heats the reaction mix to 95-98 °C to denature the

template DNA, separating it into two single strands. Depending on your sample, this usually

takes 2-5 minutes during the first thermal cycle, and 10-60 seconds for subsequent cycles.

• Annealing: When the temperature is lowered, the primers anneal to the sequences flanking

the template DNA region of interest. Depending on the sequence and melting temperature of

your primers, this step usually takes 30-60 seconds, and the optimal annealing temperature

typically lies between 45 and 60 °C.

• Extension: The temperature is increased to 72 °C, which is the ideal working temperature

for the Taq-polymerase. Depending on the synthesis rate of your polymerase, and the length

of the target sequence, it usually takes 20-60 seconds to create complementary copies

of the DNA sequence of interest.6 After approximately 25-40 cycles – depending on the

amount of DNA present at the start, and the number of amplicon copies needed for post-PCR

applications7 – the last extension step should be extended to 5 minutes or longer, allowing the

Taq-polymerase to finish the synthesis of uncompleted amplicons.5 If you can't immediately

take your samples out of the thermal cycler after the final extension step because you're busy

with other experiments, program it to cool your samples to 4 °C. For overnight runs where you

CHAPTER 1: What you need to know about PCR

5

leave your samples in the thermal cycler for hours after the final extension step, you should

opt for a holding temperature of 10 °C instead of 4 °C, as it causes less wear and tear on your

machine.

As shown in the image above, the amount of PCR product theoretically doubles at every

thermal cycle, leading to an exponential increase of PCR product. However, in reality, the phase

of exponential amplification eventually levels off and reaches a plateau because the reagents

have been consumed and the DNA polymerase activity decreases.

The different types of PCR

After performing a standard PCR reaction, you can determine the concentration, yield and

purity of the amplified DNA sequences using gel electrophoresis, spectrophotometry or

fluorometry. However, you can’t determine the amount of template DNA present in a sample

before amplification using standard PCR. If this is a requirement for your experiment, you have

to perform a qPCR reaction.

qPCR

qPCR – also called real-time PCR, quantitative PCR or quantitative real-time PCR – is a

technique used to detect and measure the amplification of target DNA as it is produced.

In contrast to conventional PCR reactions, qPCR requires a fluorescent intercalating dye

or fluorescently-labeled probes, and a thermal cycler that can measure fluorescence and

calculate the cycle threshold (Ct) value. Typically, the fluorescence intensity increases

proportionately to the concentration of the PCR product being formed, measuring quantities

of the target in real time.

CHAPTER 1: What you need to know about PCR

6

qPCR can be divided into dye-based methods (e.g. SYBR® Green) and probe-based methods

(e.g. TaqMan®).

RT-PCR and RT-qPCR

Another limitation of standard PCR is that it can only be used to amplify DNA sequences. If you

want to amplify RNA target sequences, you have to use RT-PCR.

RT-PCR

vPCR

Reverse transcription PCR (RT-PCR) is used to amplify RNA target sequences, such as

messenger RNA or RNA virus genomes. This type of PCR involves an initial incubation of

the RNA samples with a reverse transcriptase enzyme and a DNA primer – comprising

sequence-specific oligo (dT)s or random hexamers – prior to the PCR amplification.

For viability PCR (vPCR), each sample needs to be split into two aliquots. One aliquot is

incubated with a photoreactive intercalating dye that is unable to diffuse through intact cell

membranes of live cells. This means that it only intercalates into the DNA of dead cells. When

this aliquot is subsequently treated with a blue light, the dye binds irreversibly to the DNA. Both

aliquots are then subject to DNA purification and qPCR amplification. If they exhibit similar

qPCR signals, the target microorganisms in the sample are mostly viable. If the dye-treated

aliquot exhibits a weaker signal, the target microorganisms are mostly dead. vPCR is an

important technique in diagnostics, agriculture and food safety.

You can also perform a qPCR reaction instead of executing a standard PCR reaction after

the reverse transcription step, which produces cDNA from RNA. This PCR variant is called

RT-qPCR.

vPCR

The third limitation of standard PCR is that it cannot distinguish between the DNA of viable

and non-viable cells. You should use vPCR if this is important to your application, for example,

because you want to know if the infectious microorganisms in a clinical sample are dead or

alive.

CHAPTER 1: What you need to know about PCR

7

ddPCR

Digital droplet PCR (ddPCR) is another relatively new type of PCR. It uses fluorescently labeled

probes to detect DNA sequences of interest, and a water-oil emulsion system to split each

sample into about 20,000 nanoliter-sized droplets. After amplification, every droplet of the

sample is analyzed on its own. Droplets that contain at least one DNA sequence of interest emit

a fluorescent signal – and are consequently positive – while droplets without the DNA sequence

of interest don't fluoresce, and are therefore negative. Using the Poisson distribution, you can

then determine the concentration of the DNA sequence of interest in the original sample by

analyzing the ratio of positive to negative droplets for absolute quantification.8

An advantage of ddPCR compared to qPCR is that it's more precise. While qPCR can detect

two-fold differences in DNA target sequence variation, e.g. discriminate 1 copy from 2 copies

of a gene, ddPCR can discriminate 7 copies from 8 copies, which means that it can detect

differences as small as 1.2-fold.9 On top of that, ddPCR is better suited for multiplexing assays

if you want to determine the ratio of low abundance to high abundance DNA sequences of

interest, such as rare mutations on wild type backgrounds. When using qPCR, the fluorescent

signal from the high abundance sequences can dominate and obscure the signal from the

low abundance sequences. This risk is ruled out with ddPCR, as each droplet behaves as

an individual PCR reaction and contains either zero, one or, at most, a few sequences of

interest.10,11

ddPCR

CHAPTER 1: What you need to know about PCR

8

Due to these advantages, ddPCR is often preferred over qPCR for the detection of mutations

and SNPs (single nucleotide polymorphisms), allelic discrimination, gene expression studies,

and the analysis of copy number variations.12

Hot start PCR

If your PCR reaction results in non-specific amplification, you can try to increase the reaction

specificity using a hot start polymerase. This enzyme remains inactive during master mix

preparation and sample addition at room temperature, eliminating the risk that unintended

products and primer dimers are formed during PCR set-up.13

Nested and semi-nested PCR

Nested or semi-nested PCR are alternatives to hot start PCR that increase reaction specificity.

Nested PCR uses two sets of primers and two successive PCR reactions. The first set of

primers is designed to amplify a DNA sequence slightly longer than the sequence of interest.

During the second PCR reaction, the second set of primers that is specific to the sequence of

interest anneals to the amplicons of the first PCR reaction and helps to amplify the sequence of

interest.14,15

Nested PCR

CHAPTER 1: What you need to know about PCR

9

Semi Nested PCR

Semi-nested PCR works similarly to nested PCR. During the first PCR reaction, one primer

anneals to the sequence of interest and the second primer to a region flanking the sequence of

interest. This primer is then replaced with a second primer annealing to the region of interest

during the second PCR reaction.

The idea behind nested and semi-nested PCR is that, if non-specific products were amplified

during the first PCR reaction, these will not be amplified during the second PCR reaction, as the

primers cannot anneal to them.

Touchdown PCR

A third type of PCR developed to increase reaction specificity is touchdown PCR. The assay

set-up for touchdown PCR is identical to the set-up for standard PCR. The only difference lies in

the annealing step. During the first thermal cycle, the annealing temperature should be several

degrees above the optimal primer annealing temperature, then be lowered by 1-2 °C every

second cycle.16 These high temperatures during the first cycles avoid PCR primers forming

primer-dimers or binding to regions outside the DNA sequence of interest. The downside is

that the PCR primers don't all sufficiently bind to the template DNA, which leads to low yields.17

However, this can be tolerated, as the low yield of specific amplicons is then exponentially

amplified with every thermal cycle that is performed at the optimal annealing temperature.

CHAPTER 1: What you need to know about PCR

10

The history of PCR

As we've shown, there are many different types of PCR, and some of them have only recently

been developed. However, the foundation for PCR was laid in the 1950s:

• In 1953, James Watson and Francis Crick discovered the double-helix structure of DNA, and

suggested that there might be a possible copying mechanism for DNA.

• Four years later, Arthur Kornberg identified the first DNA polymerase that was able to copy the

template DNA, although only in one direction.

• In 1971, Gobind Khorana and his team started to work on DNA repair synthesis. Their

technique used DNA polymerase repeatedly, but employed only a single primer template

complex, which did not allow exponential amplification.

• At the same time, Kjell Kleppe from Khorana's lab proposed a two primer system that would

double the amount of DNA in a sample, but no one actually conducted the experiment to

find out whether it worked. The reason for this was probably that there was not yet a DNA

polymerase that could withstand the high temperatures of the denaturation step. This means

that they would have had to add a fresh dose of enzyme after every thermal cycle.

• In 1983, Kary Mullis, working at Cetus Corporation, added a second primer to the opposite

strand, and realized that repeated use of DNA polymerase triggers a chain reaction that will

amplify a specific DNA sequence, thus inventing PCR. The patent got approved in 1987, and

he won the Nobel Prize in Chemistry six years later.

• In 1976, the thermostable enzyme Taq-polymerase – which is typically used in PCR today

– was first isolated from the bacterium Thermus aquaticus, which had been discovered in a

hot spring of Yellowstone National Park in 1969. When it was finally incorporated into PCR

workflows in 1988, it removed the need to add a new dose of enzyme after every thermal

cycle, paving the way for the invention of automated thermal cyclers.18,19,20

CHAPTER 1: What you need to know about PCR

11

PCR troubleshooting

One of the most important troubleshooting mechanisms is to always include positive and

negative control samples.

If the sequence of interest wasn't amplified in your positive control sample, your master mix,

template DNA or thermal cycler could be the source of the problem:

• Master mix: Have you added the right volume and concentration of each reagent, and have

you cooled your reagents during master mix preparation?

• Template DNA: Have you run an agarose gel to ensure that your template DNA isn't

degraded? Is your template DNA pure enough and, if not, have you purified it?

• Thermal cycler: Is the number of thermal cycles sufficient for your assay? Have you

programmed the device correctly, and is it calibrated to ensure that it performs the reaction

steps at the right temperatures?

If the sequence of interest was amplified in your negative control sample, one or more

components of your master mix is contaminated. PCR reactions are very sensitive, and create

large number of copies of DNA sequences from minute amounts of starting material, so

contamination is a common issue. To prevent it, the right lab set-up is crucial.

CHAPTER 1: What you need to know about PCR

12

Lab set-up

Ideally, your PCR lab should have two rooms, each divided into two areas. The first room should

be exclusively used for pre-PCR activities, and divided into a master mix preparation area and a

sample preparation area. The second room should have a dedicated area for amplification, and

another one for product analysis.

If you’re lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible. In addition to the spatial separation, you could also consider setting up your PCR

reactions in the morning, and performing the amplification and analysis steps in the afternoon.

Temporally separating the different steps of your PCR reactions may limit your flexibility and

make you lose some time, but lowers the risk of aerosols with high DNA concentrations from the

analysis area contaminating your master mix and samples in the pre-PCR area.

On top of these precautionary measures, you should always work in biosafety cabinets or

laminar flow hoods when setting up your PCR reactions, use different sets of pipettes for master

mix preparation, sample preparation and analysis, and make sure that you use filter tips and

consumables that are free of DNase, RNase and PCR inhibitors.

Specificity

Another major PCR challenge is specificity. As explained before, it can be improved using hot

start, nested, semi-nested or touchdown PCR. A further option to prevent the amplification of

regions outside the DNA sequence of interest, as well as the formation of secondary structures,

is to redesign your primers.

CHAPTER 1: What you need to know about PCR

13

Use this checklist to see whether your primers meet all the requirements:

• Are your primers between 18 and 24 bp long?

• Is your target sequence length between 100 and 3000 bp for standard PCR assays, or 75

and 150 bp for qPCR assays?

• Do your primers have melting temperatures between 50 and 60 °C, and within 5 °C of

each other?

• Have you performed a gradient PCR to determine the optimal annealing temperature?

• Does the GC content of your primers lie between 40 and 60 %?

• Have you avoided runs or repeats of four or more bases or dinucleotides?

• Have you made sure that your primers are not homologous to a template DNA sequence

outside the region of interest?

• Have you checked that your primers can't form stable secondary structures?

PCR equipment

The most important PCR instrument is certainly the thermal cycler but, as the right pipetting

devices can help create faster and more efficient workflows with fewer errors, we'll also look at

a few different liquid handling options in this section.

Thermal cyclers

Before the development of thermal cyclers, scientists had to manually move their samples

between water baths of different temperatures. The first thermal cycler prototype called 'Mr.

Cycle' also used water baths to heat and cool the samples, and was developed by engineers

at Cetus Corporation, where Kary Mullis worked when he invented PCR.21 Today's instruments

work with electric heating and refrigeration units, and many different models with various

additional features are available.

For standard PCR, a thermal cycler that can heat and cool your samples to the required

temperatures might be sufficient to complete the different reaction steps. However, your

thermal cycler will need additional properties – such as gradient capability or an integrated

fluorometer – if you want to perform gradient PCR assays to optimize primer annealing

temperatures, or qPCR assays to determine the amount of template DNA present in a sample

before amplification.

CHAPTER 1: What you need to know about PCR

14

Pipettes

While the thermal cycler is the star of PCR labs, the right pipettes help you to process more

samples in less time, while ensuring maximal accuracy and precision. Electronic pipettes

offering a Repeat Dispense mode, for example, are a great option to boost the efficiency of

aliquoting master mix into an entire well plate. Adjustable tip spacing pipettes, paired with low

dead volume reagent reservoirs, can be a useful alternative to single channel pipettes when

transferring reagents and samples between different labware formats. And, if you want to

significantly cut your PCR set-up and purification time, pipetting robots or 96 and 384 channel

pipettes might be the right tool for you.

Conclusion

We hope that this article has been useful in helping you understand the mechanisms behind

the different types of PCR, and has shown you different ways to avoid contamination and nonspecific

amplification.

CHAPTER 1: What you need to know about PCR

15

1.2

Since the outbreak of the COVID-19 pandemic, PCR is on everybody's lips. However, only

people working in the lab know how difficult it can be to get the desired results using this wellestablished

technique. Out of this frustration came the popular joke that PCR should stand for

’pipette, cry, repeat’. To ensure that this stays a joke from now on, and that your PCR reactions

never drive you to despair again, we have compiled the most important tips and tricks for a

successful PCR set-up.

What is PCR?

The polymerase chain reaction (PCR) is used to amplify specific DNA sequences for

downstream use. Its inventor Kary Mullis, whose patent on PCR was approved in 1987, was

awarded the Nobel Prize in Chemistry six years later,1 and since this time, PCR has remained

one of the most essential molecular biology techniques. Genetic engineering, genotyping,

sequencing and the identification of familial relationships, to name a few examples, wouldn't be

possible without it.

PCR tips and tricks

To perform PCR reactions, you need to prepare a master mix, add template DNA, and amplify

the sequence of interest using a thermal cycler. This might seem straightforward, but it is far

from it. Calculating the required amounts of master mix reagents correctly to get the right

volume, at the right concentration, is the first challenge.

Once this is accomplished, the reagents need to be mixed together. The difficulty here is that

the liquids usually have to be cooled and they are often highly viscous, sticky and needed

in minimal quantities. In addition, work must be performed in a concentrated manner, as

distractions or interruptions can quickly lead to a situation where you no longer know which

reagents have already been added to the master mix. Errors such as skipping a tube or well can

CHAPTER 1: What you need to know about PCR

Simple PCR tips that can make or break your success

16

also easily occur when filling PCR strips or plates with master mix and adding template DNA,

especially when using single channel pipettes.

The last and probably biggest challenge is to keep your PCR reactions free from contamination.

PCR is a very sensitive assay that can create a large number of nucleic acid copies from a tiny

amount of starting material, so amplicon and sample contamination can be a huge problem.

Master mix calculations

Let's first have a look at the mathematical calculations needed to set up a PCR master mix.

We'll assume that you want to set up several PCR reactions with a volume of 50 μl each.

To calculate the required volume for each reagent, it is best to create a table (see Table 1) with

the necessary components, and fill in the stock concentrations and desired final concentrations

for the buffer, the MgCl2, the dNTPs and the primers. Then, calculate the dilution factors by

dividing the stock concentration by the final concentration. To determine the volume needed for

a single PCR reaction, divide the desired reaction volume by the dilution factor.2

For the polymerase, a slightly different equation is needed. The manufacturer of the enzyme

will tell you the amount of polymerase in one μl, e.g. 5 Units/μl. Fill in this value in the column

for the stock concentration and put the desired amount – e.g. 1.25 Units – in the column for

the final concentration. The volume needed can then be calculated as follows: 1.25 Units x

(1 μl / 5 Units) = 0.25 μl.3

The template DNA volume required depends on your sample type. You should add about 1 pg

to 10 ng of plasmid or viral DNA, and 1 ng to 1 μg of genomic DNA. In the example below, we

calculated how much you would need to use for 0.5 μl of a 1 μg/μl template DNA.4

Finally, add the required volumes for all the reagents. The difference between the desired total

reaction volume (50 μl) and the result obtained gives you the volume of PCR-grade water.5

REAGENT STOCK CONC. FINAL CONC.

(CF)

DILUTION

FACTOR

(= STOCK

CON. / CF)

VOLUME NEEDED

(= 50 ΜL / DIL.

FACTOR)

Buffer 10X 1X 10 5 μl

MgCl2 25 mM 1.5 mM 16.66 3 μl

dNTPs 10 mM 0.2 mM 50 1 μl

Forward primer 10 μM 250 nM 40 1.25 μl

Reverse primer 10 μM 250 nM 40 1.25 μl

Polymerase 5 Units/μl 1.25 Units - 0.25 μl

Template DNA 1 μg/μl - - 0.5 μl

PCR-grade water - - - 37.75 μl

Table 1: Example of a PCR master mix table

CHAPTER 1: What you need to know about PCR

17

After determining the required reagent volumes for one PCR reaction, you can simply multiply

them by your sample number (plus the negative and positive controls) to get the total volumes

for the entire PCR set-up. We recommend adding one additional aliquot to that result, as some

of the master mix may be lost during pipetting due to evaporation, adherence to the tip, or

pipetting inaccuracies.

That's it, you are now ready to set up your PCR reactions by following the best pipetting

practices listed below.

Best PCR pipetting practices

Start by preparing your master mix from all the components listed above, except the template

DNA. The huge advantage of preparing the entire quantity of master mix needed for an

experiment, and subsequently transferring single aliquots into PCR strips or plates, is that

you can pipette higher volumes with better accuracy. On top of that, it reduces pipetting steps,

making the entire process less tiring and error prone. Since pipetting mistakes cannot be

completely ruled out, you should add the master mix components in order of their price, starting

with the most affordable reagent. This way, you waste less money if you have to start over.6

Once your master mix is finished, well mixed and dispensed into tubes or plates, you can

add the template DNA. As the DNA samples are usually highly viscous and needed in small

quantities, you should either dispense them into the master mix or onto the wall of the tube or

well. After dispensing, keep the plunger depressed while dragging the tip gently along the wall

of your labware to remove any residual liquid. In addition, we recommend using low retention

tips.

If you're not using a hot start polymerase, cool your reagents throughout the entire process of

master mix preparation and sample addition, to prevent non-specific amplification.

When you are ready to load your samples into the thermal cycler, check that they are tightly

capped or sealed, and spin them down to ensure that no droplets remain on the labware wall

during amplification.

Pipetting solutions for PCR reactions

Before discussing various pipetting solutions, we would like to address one of the most

important aspects of liquid handling. No matter which pipettes you choose, ensure that they are

well maintained by regularly calibrating them and checking their performance in between uses.

The most affordable pipettes for master mix preparation would be manual single channel

models. However, as you need to accurately measure and mix several very expensive

reagents, we recommend investing in electronic single channel pipettes. The motor-controlled

piston movement guarantees that they always dispense the exact desired volume, minimizing

variability to increase the precision and accuracy of pipetting.

For the container, you can either prepare the master mix in a tube or, if you intend to transfer

it with an electronic multichannel pipette, in a low dead volume reagent reservoir. The

CHAPTER 1: What you need to know about PCR

18

ASSIST PLUS pipetting robot transferring master mix into a 384 well PCR plate

combination of an electronic multichannel pipette and a reservoir is ideal for this step, because

you can fill several tubes or wells simultaneously. On top of that, electronic multichannel

pipettes usually feature a Repeat Dispense mode, allowing you to aspirate a large volume

of master mix, then dispense it into multiple smaller aliquots. It is also possible to use an

electronic single channel pipette if you have a low throughput.

To add template DNA to the master mix aliquots, an adjustable tip spacing pipette can be

very handy if the labware format of your samples doesn't match the container used for PCR

amplification. For example, it allows you to transfer several template DNA samples from

microcentrifuge tubes to an entire row or column of a 96 well plate in one step.

High throughput labs might even want to take advantage of automated solutions for master

mix plating and sample transfer, such as a pipetting platform that is capable of automating

electronic pipettes.

CHAPTER 1: What you need to know about PCR

19

How to prevent PCR contamination

Several preventative measures should be taken to avoid contaminating your master mix or

template DNA with amplicons that were generated during previous PCRs.

One of the most effective means is to physically separate the master mix preparation, template

DNA addition, amplification and analysis areas from one another, and to work in laminar flow

or biosafety cabinets. Each work zone, and its corresponding equipment, should be cleaned

before and after an experiment, and tools used in one area should never enter another one.

When it comes to consumables, make sure you purchase sterile products that are certified to

be free from DNase, RNase and PCR inhibitors. Pipette tips should form a perfect seal with

the pipette to eliminate contamination that may occur when tips drip or fall off. Using filter tips

will also avoid the risk of aerosols entering your pipettes and contaminating subsequent PCR

reactions.

As you're a potential source of contamination too, always wear gloves to prevent introducing

enzymes, microbes and skin cells to the reaction, and change them when going from one area

to another. On top of that, keep your tubes closed whenever possible during the entire PCR

set-up.

Despite these preventative measures, you can't completely eliminate the possibility of

contaminated PCR reactions. To avoid having to throw away your entire stock of a certain

reagent if this occurs, prepare single use aliquots of your master mix components. You can also

prepare aliquots of positive and negative controls, as well as serial dilutions of standards for

quantitative PCR (qPCR) assays, ahead of time. Electronic pipettes with repeat dispense and

serial dilute modes can be helpful for this task, not only to reduce the risk of contamination, but

also to increase the efficiency of PCR set-up.

Conclusion

PCR is a fundamental technique in research, diagnostics and forensics. It often involves

pipetting minuscule reagent volumes with tricky properties, so it can be difficult to obtain the

desired results. On top of that, contamination can have a huge impact on results, as it's a very

sensitive assay. We hope that the tips and tricks provided in this article will help you make

your future PCR reactions a success. Many of these recommendations can also be applied to

other amplification assays, such as reverse transcription and qPCR, loop-mediated isothermal

amplification (LAMP) and helicase-dependent amplification (HDA).

CHAPTER 1: What you need to know about PCR

20

1.3 Setting up a PCR lab from scratch

PCR reactions are very sensitive and create a large number of copies of nucleic acids

from minute amounts of starting material. This makes them a fundamental and highly

effective molecular biology technique. However, because it is prone to amplicon and sample

contamination, planning and designing of your PCR lab space will need careful consideration.

CHAPTER 1: What you need to know about PCR

Designing your PCR lab

Ideally, a PCR lab should have two rooms with two areas, each designed for specific tasks.

The first room should be exclusively used for pre-PCR activities and divided into a master mix

preparation area and a sample preparation area. Air pressure should be slightly positive to

prevent aerosols from flowing in.

The second room should have a dedicated area for nucleic acid amplification, and another one

for product analysis. Air pressure should be slightly negative to ensure that amplicon aerosols

don't leave the room.

If you're lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible.

Having pre-PCR activities spatially separated from the amplification and analysis area – either

in different rooms or in separate benches – is very important, because you usually have a

low amount of the nucleic acid sample during preparation and a very high concentration after

CHAPTER 1: What you need to know about PCR 21

amplification. This means that if you analyze your PCR in the same space as you prepare your

master mix and samples, you may get false-positive results due to amplicon contamination.

You should also ensure that your lab set-up follows a unidirectional workflow. No materials or

reagents used in the amplification and analysis areas should ever be taken into the pre-PCR

space without a thorough decontamination. This means that you'll need dedicated equipment

for each area, e.g., two different sets of pipettes. This unidirectional workflow should also apply

to lab staff. If you've been working in the amplification and analysis areas, and you need to go

back to the pre-PCR area, change your personal protective equipment, as it may have been

contaminated by amplicon aerosols.

Another precautionary measure to take into account when setting up your PCR lab, in addition

to the spatial separation, is temporal separation. You could, for example, consider setting up

your PCR reactions in the morning, and perform the amplification and analysis steps in the

afternoon. This may limit your flexibility, but will prevent contamination issues and having to

repeat your experiment.

PCR equipment tips

PCR labs typically require a variety of equipment, such as centrifuges, vortex mixers, pipettes,

fridges and freezers, thermal cyclers and analysis instruments (e.g., electrophoresis systems).

Depending on the size of your lab and your applications, the amount of equipment you’ll need

may vary. Instead of providing you a 'shopping list', we will outline what you should look for

when purchasing equipment and consumables in order to keep contamination of your PCR

reactions to a minimum.

22 CHAPTER 1: What you need to know about PCR

Laminar flow or biosafety cabinet

Since you can never be 100 % certain that there are no amplicon aerosols in your pre-PCR

space, you should set up your PCR reactions in a laminar flow hood or biosafety cabinet,

decontaminated with a bleach solution prior to starting and after you finish your work.

Pipette tips and other consumables

Despite being more expensive than normal pipette tips, using filter tips for your PCR set-up will

avoid aerosols entering and contaminating your pipette, and avoid aerosols that might already

be present in your pipette contaminating your master mix or samples. To minimize your filter tip

consumption, first fill all your tubes with the master mix using only one tip or set of tips – if you're

using multichannel pipettes – and follow with your samples, using one tip per sample. Adding

the sample last is also recommended because it's easier to dispense it into a liquid than into an

empty tube, and because it reduces the risk of aerosolizing your sample as you pipette.

For consumables, you should make sure that you have enough small vials available in your lab

when your PCR reagents arrive. Aliquoting them into smaller containers will increase their shelf

life and prevent them from going through too many freeze/thaw rounds. If your reagents get

contaminated, it will also save you from throwing away your entire supply, as you’ll have clean

aliquots available for a second PCR.

Finally, you’ll need to make sure that all consumables and equipment are free of DNase, RNase

and PCR inhibitors. Always choose sterile products from manufacturers that can certify that

their tips and consumables are free of any of these potential contaminants.

CHAPTER 1: What you need to know about PCR 23

Cleaning and contamination control

You won’t need to worry about cleaning or contamination control when setting up your lab, but

you will when your lab is up and running. We will briefly address this topic below.

Whether you decide to set up your PCR reactions in a laminar flow hood, a biosafety cabinet

or an open bench, you will need to decontaminate your work space before and after set-up by

wiping it with a freshly made bleach solution and distilled water. The same process should be

performed in the amplification and analysis areas. You should also make sure you clean your

pipettes, equipment, doorknobs, and the handles of your fridges and freezers regularly.

Because PCR assays are so sensitive, all the preventative measures described here may still

not guarantee that your experiments will never get contaminated. It is therefore necessary

to include the appropriate controls to detect contamination early. Always include negative

and positive controls, as this will help identifying master mix contaminations, and confirm the

performance of the extraction protocol, reagents and amplification steps. Additionally, you

should monitor the positivity rate in your lab, and ensure that unexpected increases in detection

have identifiable causes, e.g., a seasonal outbreak.

Conclusion

In this article, we covered how to set up your PCR lab to ensure spatial and temporal

separation, and prevent contamination. We also outlined the key factors to consider when

purchasing equipment and consumables for your lab, to maintain safety and reduce wastage.

Lastly, we highlighted the importance of regular workspace cleaning and the use of appropriate

controls to detect any contamination early on. We hope that you are still just as excited about

setting up your PCR lab, and that this article has made the task less daunting for you.

24 CHAPTER 1: What you need to know about PCR

1.4 qPCR: How SYBR® Green and TaqMan®

real-time PCR assays work

qPCR, or real-time PCR, is a widely used method to quantify DNA sequences in samples.

This article gives you a comprehensive introduction to the topic, explaining how dye-based

and probe-based qPCR assays (like SYBR Green and TaqMan) work, how to validate your

amplification experiments, and how to analyze your qPCR data.

qPCR vs PCR vs RT-PCR

Before explaining how qPCR works, we would like to briefly outline its difference from standard

PCR and RT-PCR.

Whereas standard PCR monitors DNA amplification upon reaction completion, qPCR uses

fluorescent signals to monitor DNA amplification as the reaction progresses. This is why qPCR

is also referred to as real-time PCR, quantitative PCR or quantitative real-time PCR.

RT-PCR, not to be confused with real-time PCR, stands for reverse transcription PCR and can

be used to amplify RNA target sequences. It involves an initial incubation of the sample RNA

with a reverse transcriptase enzyme and a DNA primer before amplification.

How qPCR works

qPCR relies on fluorescence from intercalating dyes or hydrolysis probes to measure DNA

amplification after each thermal cycle. The most common dye-based method is SYBR Green,

and the most common probe-based method is TaqMan, which is why this article will focus on

these two qPCR techniques.

CHAPTER 1: What you need to know about PCR 25

SYBR Green qPCR

Like standard PCR, the SYBR Green protocol consists of denaturation, annealing and

extension phases. The difference being that you add some double-stranded DNA binding dye,

SYBR Green I, to your master mix during qPCR setup. This fluorescent dye intercalates into

double-stranded DNA sequences during the extension phase, where it shows a strong increase

in fluorescent signal. Measuring this signal at the end of every thermal cycle will allow you to

determine the quantity of double-stranded DNA present.

The downside of the SYBR Green assay is that the dye binds to any double-stranded DNA

sequence. This means that you could also detect fluorescence emitted from non-specific

qPCR products, such as primer dimers. To eliminate this risk, check the reaction specificity by

performing a melting curve analysis, explained later in the article, or use the TaqMan method.

TaqMan qPCR

Instead of using intercalating dyes, this assay uses TaqMan probes with a 5' fluorescent

reporter dye and a 3' quencher dye. These probes are target-specific, and only bind to the

DNA sequence of interest downstream of one of the primers during the annealing step. When

the enzyme Taq-polymerase encounters the TaqMan probe during the extension phase, it

displaces and cleaves the 5' reporter dye. Once the reporter dye has been separated from

the quencher dye, its measurable fluorescent signal at the end of every qPCR cycle increases

significantly. The second DNA strand is synthesized in parallel but, as no probe is attached to it,

this process can't be monitored by fluorescence measurements.

Compared to the SYBR Green assay, the use of TaqMan probes is more expensive, but also

offers two significant advantages:

• the TaqMan assay only measures amplification progression of the target sequence, as the

probes are target specific.

• you can monitor the quantity of various qPCR products in a single reaction by adding different

primers and TaqMan probes with different reporter dyes to the master mix. This multiplex

approach allows you to detect several fluorescent signals at the end of every thermal cycle.

26 CHAPTER 1: What you need to know about PCR

Amplification plot

For both qPCR methods, data is visualized in an amplification plot, with the number of thermal

cycles on the x-axis, and the fluorescent signals detected on the y-axis:

CHAPTER 1: What you need to know about PCR 27

As can be seen, fluorescence remains at background levels during the first thermal cycles.

Eventually, the fluorescent signal reaches the fluorescence threshold, where it is detectable over

the background fluorescence. The cycle number at which this happens is called the threshold

cycle (Ct). If the Ct value for a sample is high, it means that little starting material was present,

and vice versa. Please note that you should always analyze at least three replicates of each

sample, as tiny pipetting errors during qPCR set-up can result in huge differences in Ct values.

The Ct value is sometimes also referred to as crossing point (Cp), take-off point (TOP) or

quantification cycle (Cq) value, with MIQE guidelines suggesting using Cp value to standardize

terminology.1 In this article we will continue to call it Ct, as this is the most commonly used term.

MIQE guidelines

The MIQE (Minimum Information for Publication of Quantitative Real-Time PCR Experiments)

guidelines describe the minimum information necessary for evaluating qPCR experiments.

When publishing a manuscript, the scientist needs to provide all relevant experimental

conditions and assay characteristics described by the MIQE guidelines, allowing reviewers

to assess the validity of the protocols used, and enabling other scientists to reproduce the

experiments.

Validation of qPCR assays

qPCR amplification plots can be analyzed using absolute or relative quantification. However,

before explaining qPCR data analysis, we need to quickly discuss how to determine reaction

efficiency and specificity. You don't need to perform these steps after every qPCR experiment,

but should always validate these two values when setting up a new qPCR protocol or changing

your current workflows.

Reaction efficiency

A perfect qPCR assay would have a reaction efficiency of 100 %, which means that the number

of template DNA copies would double at every thermal cycle. As this is almost impossible to

achieve in practice, reaction efficiencies between 90 and 110 % are considered to be ideal.

To calculate the reaction efficiency of your assay, you need to set up a 10-fold serial dilution

of an undiluted sample with a known amount of template DNA. After running a qPCR, create a

standard curve with the log of the starting quantity on the x-axis and the Ct values on the y-axis.

28

Using the equation for the linear regression line (y = mx + b), you can now determine the

reaction efficiency as follows2:

Efficiency = (10(-1/m)-1) x 100

In our example, m would be -3.5826, resulting in a reaction efficiency of 90.1634 %.

Reaction specificity

Reaction specificity can be determined using a melting curve analysis, allowing you to identify

non-specific qPCR products and primer-dimers. To perform a melting curve analysis, run a

qPCR assay with a fluorescent intercalating dye like SYBR Green I. After amplification, the

thermal cycler increases the temperature step by step while monitoring fluorescence. As

the temperature increases, the dsDNA qPCR products present will denature, resulting in a

decreasing fluorescent signal:

CHAPTER 1: What you need to know about PCR

CHAPTER 1: What you need to know about PCR 29

Then, plot the change in slope of this curve as a function of temperature to obtain a melting

curve:

If you're observing only one melting peak like the image above, your qPCR assay is specific.

If there are several melting peaks, primer-dimers and/or non-specific products were amplified

during qPCR, and you should redesign your experiment to increase its specificity.

30 CHAPTER 1: What you need to know about PCR

Analysis of qPCR data

qPCR data can be analyzed by absolute or relative quantification, and the method suitable

for your experiment depends on your goal. Absolute quantification allows you to determine

the quantity of starting material that was present in a given sample before amplification. For

example, this method can be used to determine the viral load of a patient sample. Relative

quantification is applied to compare levels or changes in gene expression between different

samples. For example, it is helpful to investigate whether the expression of a certain gene is

higher in a tumor sample than in a healthy control sample.

Absolute quantification

After qPCR amplification, you will have produced an amplification plot, and know the Ct value

of each sample. To find the quantity of starting material present in your samples, you need to

compare these values to a standard curve. As seen above in the section on reaction efficiency,

a standard curve is obtained by amplifying a serial dilution of a sample with a known amount of

template DNA, then plotting the Ct values against the log of the starting quantities.

The equation for the linear regression line of the standard curve (y = mx + b) will then allow you

to calculate the quantity of starting material for each sample. As y corresponds to the Ct value,

and x to the log quantity, the equation for the linear regression line is equivalent to:

Ct = m(log quantity) + b

Solving this equation for the quantity will give you the formula:

Quantity = 10((Ct-b)/m)

This will allow you to quickly determine the quantity of starting material in each sample.

Y = mx + b → Ct = m(log quantity) + b → Quantity = 10((Ct-b)/m)

Relative quantification

To compare levels or changes in target gene expression between different samples and a

control sample, you first need to define a reference gene whose expression is unregulated.

Then, run a qPCR to obtain the Ct values for the reference gene, target gene in your samples,

and the control sample.

If the reaction or primer efficiencies for the reference and target genes are near 100 %, and

within 5 % of each other, you can then use the ΔΔCt method – also called the Livak method – to

determine the expression rate of the target gene in your samples. However, if the efficiencies

are further apart, you should use the Pfaffl method. To learn how to calculate reaction

efficiencies, please refer to the 'Reaction efficiency' section earlier in the article.

CHAPTER 1: What you need to know about PCR 31

The calculations for the two methods are as follows:

ΔΔCt method

Normalize the Ct of the target gene to the Ct of the reference gene for each sample and the

control sample:

ΔCt(sample) = Ct(target gene) – Ct(reference gene)

ΔCt(control) = Ct(target gene) – Ct(reference gene)

Normalize the ΔCt of each sample to the ΔCt of the control sample:

ΔΔCt(sample) = ΔCt(sample) – ΔCt(control)

Since the calculations are in logarithm base 2, you must use the following equation to get the

normalized expression ratio for each sample:

Normalized expression ratio = 2-ΔΔCt(sample)

Pfaffl method

Calculate the ΔCt of the target gene for each sample:

ΔCt(target gene) = Ct(target gene in control) – Ct(target gene in sample)

Calculate the ΔCt of the reference gene for each sample:

ΔCt(reference gene) = Ct(reference gene in control) – Ct(reference gene in sample)

Calculate the normalized expression ratio for each sample:

Normalized expression ratio = ((Etarget gene)ΔCt(target gene)) / ((Ereference gene)ΔCt(reference gene))

Etarget gene: Reaction efficiency of the target gene

Ereference gene: Reaction efficiency of the reference gene

The normalized expression ratio obtained using the ΔΔCt or the Pfaffl method is the fold

change of the target gene in your sample relative to the control. A normalized expression ratio

of 1.2 would mean that you have a gene expression of 120 % relative to the control.

Conclusion

We hope that this article answered all your questions regarding qPCR methods, assay

validation and data analysis.

32

1.5 How to design primers for PCR

PCR is one of the most widespread molecular biology applications, yet it is anything but simple

to perform. Common issues – such as a low product yield or non-specific amplification – are

often caused by poorly designed PCR primers. We have therefore summarized the most

important information on designing PCR primers to help you overcome these challenges.

What is a PCR primer?

Primers – also called oligonucleotides or oligos – are short, single-stranded nucleic acids used

in the initiation of DNA synthesis. During PCR reactions, they anneal to the plus and minus

strands of the template DNA, flanking the sequence that needs to be amplified.

How to design PCR primers?

PCR primers have to be tailored to both the region of interest of your template DNA and your

reaction conditions. This means that, unlike the other components of the PCR master mix, you

can't just buy them, but need to design them yourself using a primer design tool. These tools

allow you to set parameters such as primer length, melting temperature, GC content and more.

Read on to learn what the optimal values for each of these parameters are, and how they affect

your PCR assay.

CHAPTER 1: What you need to know about PCR

CHAPTER 1: What you need to know about PCR 33

Primer length

The optimal length of a PCR primer lies between 18 and 24 bp. Longer primers are less efficient

during the annealing step, resulting in a lower amount of PCR product. Conversely, shorter

primers are less specific during the annealing phase, leading to more non-specific binding and

amplification. However, there are exceptions to this rule. For example, some scientists have

successfully used miniprimers that are 10 bp long to expand the scope of detectable sequences

in microbial ecology assays.

Target sequence length

The target sequence to be amplified should ideally be between 100 and 3000 bp for standard

PCR assays, and 75 and 150 bp for qPCR assays. Longer sequences usually need special

enzymes and reaction conditions to ensure that they are completely and specifically amplified.

Primer melting temperature

The primer melting temperature (Tm) can be defined as the temperature at which half of the

primers dissociate from the template DNA. It is usually between 50 and 60 °C, and the melting

temperatures of the forward and reverse primers should be within 5 °C of each other. If the two

melting temperatures are further apart, it won't be possible to find an annealing temperature

that allows both primers to bind to the template DNA.

Most primer design tools use the nearest neighbor method to calculate primer melting

temperatures, as it's the most accurate. However, if you want to make an approximate

calculation yourself, you can use this formula:

Tm = 4 °C x (G+C) + 2 °C x (A+T)

Tm: melting temperature

G, C, A, T: number of nucleobases (guanine, cytosine, adenine, thymine) in the primer

As indicated in the formula above, G-C bonds are harder to break than A-T bonds – because

G-C base pairs are linked by three hydrogen bonds, and A-T base pairs by two – and the length

of the primer also impacts its melting temperature. This means that you can either increase the

GC content of a primer (provided the template allows for this), or slightly extend its length if its

melting temperature is too low.

34

Primer annealing temperature

The primer annealing temperature (Ta) is the temperature needed for the annealing step of

the PCR reaction to allow the primers to bind to the template DNA. The theoretical annealing

temperature can be calculated as follows:

Ta = 0.3 x Tm(primer) + 0.7 x Tm(product) – 14.9

Ta: primer annealing temperature

Tm(primer): lower melting temperature of the primer pair

Tm(product): melting temperature of the PCR product

Once you've calculated the theoretical annealing temperature, the optimal annealing

temperature needs to be determined empirically. To achieve this, perform a gradient PCR,

starting a few degrees below the calculated annealing temperature, and ending a few degrees

above. After amplification, run a gel, and the sample producing the clearest band contains the

largest quantity of PCR product, making its annealing temperature the optimal one for your

primers. Usually, you'll get a value that is 5 to 10 °C lower than the primer melting temperature.

CHAPTER 1: What you need to know about PCR

35

It's important to determine the optimal annealing temperature, as primers could form hairpins or

bind to regions outside the DNA sequence of interest if it's too low, producing non-specific and

inaccurate PCR products. If the annealing temperature is too high, the primers won't sufficiently

bind to the template DNA, and you'll obtain little to zero amplicons.

CHAPTER 1: What you need to know about PCR

GC content

As seen before, G-C base pairs are stronger than A-T base pairs, which means that a higher

GC content ensures a more stable binding between the primers and the template DNA. The

optimal GC content of a primer lies between 40 and 60 %, and primers should have two to three

Gs and Cs at the 3' end to bind more specifically to the template DNA.

Runs and repeats

Avoid runs of four or more single bases – such as ACCCCC – or dinucleotide repeats (for

example, ATATATATAT), as they can cause mispriming.

Cross homology

If a primer is homologous to a template DNA sequence outside the region of interest, these

sequences will be amplified too. Therefore, you should always test the specificity of your

primer design against genetic databases; for example, by ‘blasting’ them through NCBI BLAST

software.

36

Your PCR product yield will be less if secondary structures form and remain stable above the

annealing temperature of your reaction, as the primers bind to themselves or another primer

instead of the template DNA. This is why your primer design tool should be able to check for,

and warn you of stable secondary structures.

Mismatches and degenerated positions

Mismatches are primer bases that aren't complementary to the target sequence. They can be

tolerated to a certain extent, and are sometimes even necessary; for example, when performing

a multi-template PCR to amplify a set of similar target sequences from different bacteria with

a single set of primers. Degenerate primers could help if mismatches negatively impact the

performance of your PCR.

Degenerate primers have several different nucleotides in some of their positions. For example,

instead of A you could have an equal concentration of A and T in a certain position. The codes

for the different nucleotide combinations available for degenerate primers are as follows:

CHAPTER 1: What you need to know about PCR

Secondary structures

There are three different types of secondary structures – also called primer dimers – that can

form during a PCR assay:

• Hairpins: caused by intra-primer homology – when a region of three or more bases is

complementary to another region within the same primer – or when a primer melting

temperature is lower than the annealing temperature of the reaction.

• Self-dimers: formed when two same sense primers have complementary sequences – interprimer

homology – and anneal to each other.

• Cross-dimers: formed when forward and reverse primers anneal to each other when there is

inter-primer homology.

37

IUPAC NUCLEOTIDE CODE BASE

R A or G

Y C or T

S G or C

W A or T

K G or T

M A or C

B C or G or T

D A or G or T

H A or C or T

V A or C or G

N Any base

Conclusion

This article summarized the key points to consider when designing PCR primers to help avoid

common issues like low product yield or non-specific amplification. We covered optimal primer

and target sequence lengths, and ideal primer melting and annealing temperatures. We also

provided helpful tips for other crucial factors such as GC content, runs and repeats, cross

homology and the danger of stable secondary structures. Lastly, the article highlighted the

value and pitfalls of mismatches and degenerated positions. That's it, after reading about all of

this, you are sure to be a 'PCR Primer Pro'!

CHAPTER 1: What you need to know about PCR

38

CHAPTER 2:

INTEGRA Biosciences’ PCR solutions

PCR is a robust method, but it’s comprised of numerous stages, involving multiple precise

pipetting steps that often prove time consuming and prone to errors. Temperature-sensitive

reagents and samples may affect accuracy, and the varying viscosities of samples, as well as

‘sticky’ DNA, can be difficult to handle. On top of this, the repetitive nature of this work can also

frequently result in user fatigue and handling mistakes.

Fortunately, the right tools can eliminate your pipetting predicaments, vastly improving the

reproducibility and productivity of your PCR workflows. Here, we will demonstrate how our

range of liquid handling solutions are perfect for PCR applications, allowing you to create a

faster and more efficient workflow with fewer errors.

Manual and electronic pipettes

A good starting point for lower throughput PCR applications – up to half a plate per day – is our

EVOLVE single or multichannel pipettes, which feature convenient volume adjustment dials to

increase the accuracy and speed of manual handling. Our range of VIAFLO electronic pipettes

is also suitable for low throughput PCR set-up, and can easily handle up to eight plates per day.

CHAPTER 2: INTEGRA Biosciences’ PCR solutions

Learn more

about

EVOLVE

Learn more

about

VIAFLO

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 39

Learn more

about

VOYAGER

Adjustable tip spacing pipettes

PCR set-up usually requires transferring liquids between different labware formats which is

tedious and highly error prone. Our VOYAGER adjustable tip spacing pipettes solve these

problems, increasing speed and eliminating transfer errors, while ergonomic single-handed

operation leaves the other hand free to handle labware.

96 and 384 channel pipettes

We have a wide range of options perfect for productive high throughput PCR set-up – more

than eight plates per day – which are suitable for different lab sizes and budgets. Our

VIAFLO 96 and VIAFLO 384 channel handheld electronic pipettes, as well as the

MINI 96 channel portable electronic pipette, can reduce handling steps while

increasing productivity and reproducibility.

Learn more

about

MINI 96

Learn more

about

VIAFLO 96

and VIAFLO 384

40 CHAPTER 2: INTEGRA Biosciences’ PCR solutions

Pipetting robots

INTEGRA also offers pipetting robots for high throughput laboratories, or for labs that want to

reduce the risk of contamination due to manual processing. For example, the ASSIST PLUS

pipetting robot can automate the D-ONE single channel pipetting module for master mix

preparation, and VIAFLO and VOYAGER multichannel pipettes to take care of the multiple

pipetting steps in PCR workflows.

Learn more

about

D-ONE

Learn more

about

GRIPTIPS

Learn more

about

ASSIST PLUS

Pipette tips

INTEGRA has developed GRIPTIPS pipette tips to complement its range of pipetting solutions.

GRIPTIPS are free from RNase, DNase and PCR inhibitors, and perfectly fit all INTEGRA

pipetting solutions, reducing the risk of contamination from tips that leak or fall off.

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 41

Learn more about

sample transfers

from plate to plate

Learn more about

sample transfers

from tubes to plates

Sample reformatting

The transfer of samples between different labware formats is a slow, tedious and highly

error-prone procedure when performed manually with a single channel pipette. The

combination of the ASSIST PLUS pipetting robot and VOYAGER adjustable tip spacing

pipette provide a novel solution for automated, accurate and efficient liquid transfer of multiple

samples in parallel. For even higher throughput applications, the VIAFLO 96, VIAFLO 384

and MINI 96 offer a fast solution for whole plate transfers.

42 CHAPTER 3: Application Notes

CHAPTER 3:

Application notes

Our pipetting instruments are used across a broad spectrum of life sciences applications. To

help share this knowledge and experience of using INTEGRA products with the wider scientific

community, we have developed an application database which contains a wide range of useful

application notes. Here are some of the most relevant app notes related to PCR protocols and

workflows.

3.1 Efficient and automated 384 well qPCR set-up

with the ASSIST PLUS pipetting robot

Using the ASSIST PLUS pipetting robot to automate set-up for a 384 well

plate qPCR

Setting up a qPCR is a tedious process consisting of multiple pipetting steps. One particularly

challenging task is reformatting from microcentrifuge tubes into a 384 well plate, which is time

consuming and requires a lot of concentration. Another common problem is the loss of valuable

and expensive substances, such as master mix and

precious samples, due to the reservoir dead volume.

The ASSIST PLUS pipetting robot, in combination with

the VIAFLO and VOYAGER electronic pipettes,

streamlines the workflow and increases the throughput

and the reproducibility of qPCR set-ups, with minimal

manual input. The loss of expensive substances or

valuable samples due to reformatting errors is

eliminated. The unique design of the ASSIST PLUS

pipetting robot, together with the intuitive

VIALAB software, offers

exceptional flexibility and

straightforward implementation.

CHAPTER 3: Application Notes 43

Key benefits

• Automating the qPCR set-up with the

VIAFLO 16 channel electronic pipette and

the ASSIST PLUS pipetting robot allows

considerably faster sample preparation,

freeing up time for scientists to focus on

other experiments.

• Automation of VOYAGER adjustable tip

spacing pipettes with the ASSIST PLUS

offers a reliable pipetting method that

requires minimal manual intervention and

eliminates the risk of reformatting errors.

• The use of low retention GRIPTIPS with

heightened hydrophobic properties and

SureFlo™ low dead volume reservoirs with

an anti-sealing array helps to save precious

samples and master mix. Combined with

the high pipetting accuracy and precision

of the ASSIST PLUS pipetting robot, this

enables exceptionally low dead volumes to

be achieved.

• The ASSIST PLUS pipetting robot, in

combination with the intuitive VIALAB

software, is quick to set up and easy to use.

Overview: qPCR set-up

The ASSIST PLUS pipetting robot is used to set up a 384 well format qPCR by pipetting 64

samples in triplicate with two different master mixes for the detection of two genes of interest

(GOI 1 and GOI 2).

The protocol is divided into two programs that guide the user through all the steps of the qPCR

set-up:

• Program 1: Mastermix_qPCR

• Program 2: Samples_qPCR

The ASSIST PLUS pipetting robot operates a VIAFLO 16 channel 125 μl electronic pipette with

125 μl sterile, filter, low retention GRIPTIPS for program 1 and a VOYAGER 8 channel 12.5 μl

electronic pipette with 12.5 μl sterile, filter, low retention GRIPTIPS for program 2.

44

Experimental set-up: Program 1 - master mix

transfer (Mastermix_qPCR)

Prepare the pipetting robot deck as follows (Figure 1):

Deck position A: Dual reservoir adapter – 2 x 10 ml reagent

reservoir with SureFlo anti-sealing array

(Figure 2) containing master mix 1 and 2.

Deck position B: 384 well PCR plate, placed on an INTEGRA

cooling block in the landscape position.

CHAPTER 3: Application Notes

Figure 1: Set-up for the master mix transfer. Position A: dual reservoir adapter with 2 x 10 ml reagent

reservoirs with SureFlo anti-sealing array. Position B: 384 well PCR plate, placed on an INTEGRA

cooling block.

Figure 2: The INTEGRA dual reservoir adapter accommodates both 10 ml reagent reservoirs on one

deck position.

CHAPTER 3: Application Notes 45

Step-by-step procedure

1. Transfer master mixes into the 384 well plate

Add master mixes 1 and 2 into the left and right sides of the 384 well PCR

plate, respectively.

Use an EVOLVE 5000 μl manual pipette with

5000 μl sterile, filter, low retention GRIPTIPS to

fill the left 10 ml reagent reservoir with SureFlo

anti-sealing array with 1.6 ml of master mix 1 and

the right reservoir with 1.6 ml of master mix 2

(position A). Select and run the VIALAB program

‘Mastermix_qPCR’ on the VIAFLO 16 channel

125 μl electronic pipette with 125 μl sterile, filter,

low retention GRIPTIPS. The ASSIST PLUS

pipetting robot automatically transfers 7.5 μl of

master mix 1 (pink) into the left half of the 384 well

PCR plate and 7.5 μl of master mix 2 (blue) into the

right half (Figure 3) using the Repeat Dispense

mode with a tip touch on the surface of the liquid to

increase pipetting precision. Figure 4 shows the

pipetting robot transferring the master mix into a

384 well plate.

Tips:

• Pre- and post-dispense steps are recommended

to increase the accuracy and precision of

pipetting. The pre- and post-dispense volumes

should be between 3 and 5 % of the nominal

volume of the pipette.

• The low retention GRIPTIPS are made from a

unique polypropylene blend with heightened

hydrophobic properties for superior accuracy

and precision while pipetting viscous and low

surface tension liquids.

• The reservoirs’ SureFlo anti-sealing array and

a unique surface treatment that spreads liquid

evenly enable the pipette tips to sit on the bottom

and still aspirate liquids accurately, reducing

dead volumes.

Figure 3: Pipetting scheme for master mixes 1 (pink) and 2 (blue).

Figure 4: Example of the ASSIST PLUS pipetting robot

transferring a master mix into a 384 well PCR plate.

46 CHAPTER 3: Application Notes

Experimental set-up: Program 2 - sample transfer

(Samples_qPCR)

Prepare the pipetting robot deck as follows (Figure 5):

Deck position B: 384 well PCR plate, placed on an INTEGRA

cooling block.

Deck position C: INTEGRA 1.5 ml microcentrifuge tube rack,

with tubes containing samples 1-32.

Figure 5: Set-up for the sample transfer protocol. Position B: 384 well PCR plate, placed on an INTEGRA

cooling block. Position C: INTEGRA 1.5 ml microcentrifuge tube rack, with tubes containing samples 1-32

(Figure 6).

Figure 6: Example of the ASSIST PLUS pipetting samples from the INTEGRA microcentrifuge tube rack

into a 96 well plate.

VOYAGER - 12.5 μl – 8CH

12.5 μl GRIPTIP,

sterile, filter

B 384 well PCR Sapphire on 384 well cooling block – 45 μl C Rack for 1.5 ml microcentrifuge tubes – 1500 μl

CHAPTER 3: Application Notes 47

Step-by-step procedure

1. Sample transfer into the 384 well plate

Add the 64 samples in triplicate to the master mixes.

Place samples 1-32 in an INTEGRA 1.5 ml microcentrifuge tube rack on position C. Run the

VIALAB program ‘Samples_qPCR’ on a VOYAGER 8 channel 12.5 μl electronic pipette to start

the sample transfer. The ASSIST PLUS transfers 2.5 μl of the first 32 samples in triplicate into

master mixes 1 and 2 (Figure 7, yellow/brown), using the Repeat Dispense mode with a tip

touch on the side of the well to make sure that no droplets adhere to the GRIPTIPS. After this

step, a prompt informs the user to place the second series of samples (33-64) on position C.

The ASSIST PLUS pipetting robot continues by transferring 2.5 μl of the samples in triplicate

into the other half of master mixes 1 and 2 (Figure 7, green).

Tip: Use sterile, filter, low retention GRIPTIPS for optimal liquid recovery of precious solutions,

such as the master mix and samples.

Figure 7: Pipetting scheme of the qPCR assay.

master mix 1 master mix 2

Sample

33 - 64

Sample

1 - 32

48 CHAPTER 3: Application Notes

Remarks

VIALAB software:

The VIALAB program can easily be adapted to fit the user’s demands, especially if specific

labware, incubation times or protocols are needed.

Partial plates:

The programs can be adapted at any time to a different number of samples, giving

laboratories total flexibility to meet current and future demands.

Conclusion

• The time required for a 384 well qPCR set-up can be reduced from 1.5 hours using

a single channel pipette to 12 minutes using the ASSIST PLUS pipetting robot in

combination with VIAFLO 16 channel and VOYAGER 8 channel pipettes.

• The ASSIST PLUS, together with the VOYAGER adjustable tip spacing pipette,

guarantees perfectly reproducible test results and eliminates all risks of reformatting

errors when transferring samples from microcentrifuge tubes into a 384 well plate.

• INTEGRA’s low retention GRIPTIPS increase pipetting precision for viscous or low

surface tension liquids. The reagent reservoirs with SureFlo anti-sealing array reduce the

dead volume of costly reagents and precious samples.

• The intuitive VIALAB qPCR program is quick to set up and easy to use or adapt to other

pipetting protocols.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 3: Application Notes 49

3.2 Automated RT-PCR set-up for

COVID-19 testing

How to prepare RT-PCR plates for SARS-CoV-2 detection

with the ASSIST PLUS

The emergence and outbreak of the novel coronavirus

SARS-CoV-2 (COVID-19) has placed unprecedented

demands on laboratories testing for COVID-19, leaving

scientific staff to contend with a spiraling influx of patient

samples and a rapid, continuous growth in workload.

Laboratories need additional automated liquid handling

instruments for viral nucleic acid extraction

and RT-PCR set-up – which are the

most labor-intensive processes in

the diagnostic testing workflow – to

increase the sample throughput

capacity, reduce manual

intervention by laboratory

analysts and fast track the

development of COVID-19 assays.

The ASSIST PLUS pipetting robot together with a VOYAGER

adjustable tip spacing pipette, low retention GRIPTIPS and SureFlo

10 ml reagent reservoirs were successfully used for RT-PCR set-up in

COVID-19 testing laboratories.

50 CHAPTER 3: Application Notes

Key benefits

• The full automation capability of the

ASSIST PLUS reduces manual intervention

and frees highly valuable time for laboratory

personnel in this critical COVID-19

pandemic.

• The compact and easy-to-use

ASSIST PLUS pipetting robot allows

fast set-up regarding installation and

programming, allowing labs to immediately

increase their sample processing capacity

and fast track assay development for

COVID-19 sample testing.

• VOYAGER adjustable tip spacing pipettes

in combination with the ASSIST PLUS

provide unmatched pipetting ergonomics by

automatically reformatting patient samples

from tube racks into 384 well plates.

• Optimal pipette settings, including tip

immersion depth, pipetting speeds and

angles, deliver reproducible, precise and

accurate results, with no contamination

observed in controls or patient samples.

• The use of INTEGRA’s low dead volume,

SureFlo 10 ml reagent reservoirs, together

with low retention GRIPTIPS, demonstrated

excellent results, enabling efficient handling

of the precious and expensive one-step RTPCR

master mix used for patient testing.

Overview: Automated RT-PCR set-up

The ASSIST PLUS pipetting robot is used to automate testing of suspected COVID-19

positive cases in a 384 well plate. The pipetting robot operates a VOYAGER 12 channel 50 μl

electronic pipette with 125 μl sterile, filter, low retention GRIPTIPS. To double the available

testing capacity and, concurrently, decrease the cost per test of expensive one-step RT-PCR

reagents of dwindling availability, the total PCR reaction volume was miniaturized, reducing it

to 10 μl – inclusive of 7.5 μl one-step RT-PCR master mix and 2.5 μl of nucleic acid template.

The templates were extracted from combined nasopharyngeal/oropharyngeal flocked swabs

or sputum samples. The following procedure is based on the protocol used by the Microbiology

and Molecular Pathology Department at Sullivan Nicolaides Pathology (SNP) – part of the

Sonic Healthcare Group – in Brisbane, Australia.

The protocol is divided into two parts:

• Program 1: Add the master mix (1-COVID-19)

• Program 2: Add the nucleic acid template (2-COVID-19)

CHAPTER 3: Application Notes 51

Experimental set-up: Program 1

Deck position A: 10 ml reagent reservoir with

SureFlo anti-sealing array containing

3 ml of one-step RT-PCR master mix.

Deck position C: 384 well plate placed on a PCR 384 well

cooling block, allowing the master mix and

samples to be kept cold, and enabling exact

positioning of the PCR plate on the deck.

Figure 1: The set-up for program 1-COVID-19.

VOYAGER - 50 μl – 12CH

50/125 μl GRIPTIP, sterile, filter,

low retention

A Multichannel reservoir – 10ml C PCR cooling block 384_system

52 CHAPTER 3: Application Notes

Step-by-step procedure

1. Add the master mix

Fill the 384 well plate with the one-step RT-PCR master mix.

Place the one-step RT-PCR master mix in a 10 ml sterile, polystyrene reagent reservoir with

INTEGRA’s SureFlo anti-sealing array. Set up the deck with the required labware, as indicated

in Figure 1. Select the VIALAB program 1-COVID-19. The VOYAGER pipette automatically

transfers the master mix from the reservoir into the 384 well plate (LightCycler® 480 Multiwell

Plate, Roche) using the Repeat Dispense mode with tip touch. Each well of the plate is filled

with 7.5 μl of master mix.

Tips:

• Using a 10 ml reagent reservoir with SureFlo anti-sealing array allows a very low dead

volume (<20 μl) to minimize the loss of expensive reagent of dwindling availability

(see Figure 2).

• The combination of a low pipetting speed – set at 2 – and low retention GRIPTIPS shows

excellent results when pipetting the viscous and foamy master mix.

• Pre- and post-dispense settings, together with the tip touch option, guarantee reproducible,

precise and accurate pipetting results (see Figure 2).

• The PCR cooling block is used as a support to fix the position of the 384 well plate on the

deck, ensuring exact tip positioning when pipetting. The cooling block also helps to keep

samples and reagents cool if required by the protocol.

Figure 2: Precise and accurate dispensing of one-step RT-PCR master mix from the low dead

volume reagent reservoir to the 384 well plate.

CHAPTER 3: Application Notes 53

Experimental set-up: Program 2

Deck position A and B: FluidX Cluster 0.7 ml tubes containing the

nucleic acid templates. The tubes are stored

in a 96-format rack. A total of four sample

racks are used for the protocol (two on

position A and two on position B).

Deck position C: 384 well plate placed on a PCR 384 well

cooling block.

Figure 3: The set-up for program 2-COVID-19.

2. Add the nucleic acid templates

Transfer the samples from four 96-format tube racks to the 384 well plate.

Nucleic acid templates extracted from combined nasopharyngeal/oropharyngeal flocked

swabs or sputum samples are stored in FluidX Cluster 0.7 ml tubes placed in a 96-format

rack. The VOYAGER pipette transfers 2.5 μl of template from the tubes to the 384 well plate,

automatically changing the GRIPTIP pipette tips after each dispense. Both position A and B

are used to house the samples on the deck (see Figure 3). The pipette prompts the user when

it is time to replace the tube racks on the deck. After user confirmation, the VOYAGER pipette

continues reformatting the samples from tubes to the plate.

50/125 μl GRIPTIP, sterile, filter,

low retention

VOYAGER - 50 μl – 12CH

A FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl

B FluidX 96-formal, 0.7 ml Internal Thread Tube, V-Bottom

– 700 μl C PCR cooling block 384_system

54

Tips:

• The VOYAGER pipette’s tip spacing capability combined with automatic Tip Change ensures

easy and rapid sample transfer without risk of contamination or reformatting errors.

• Using an air gap of 1.5 μl when aspirating the viral nucleic acid template eliminates the risk of

contamination risk during pipette tip travel.

Note: Automated RT-PCR testing for COVID-19 with the ASSIST PLUS can also be

performed using a VOYAGER 8 channel 50 μl electronic pipette (see Figure 5).

Figure 4: Easy and rapid transfer of patient nucleic acid templates from the tube rack to the 384 well

plate using the VOYAGER adjustable tip spacing pipette together with the ASSIST PLUS pipetting robot.

Figure 5: Automated RT-PCR testing for COVID-19 using the ASSIST PLUS pipetting robot together with

a VOYAGER 8 channel adjustable tip spacing pipette, as performed in the Microbiology and Molecular

Pathology Department at SNP.

CHAPTER 3: Application Notes

55

Remarks

4 Position Portrait Deck:

If your process allows, the protocol can be compiled into one simple program using the

4 Position Portrait Deck option on the ASSIST PLUS (see Figure 6).

96 well plates:

The protocol can be readily adapted to 96 well format.

VIALAB software:

The VIALAB programs can be easily adapted to your specific labware and protocols.

CHAPTER 3: Application Notes

Figure 6: Example set-up of the 4 Position Portrait Deck when combining programs 1-COVID-19 and

2-COVID-19 in one program.

50/125 μl GRIPTIP, sterile, filter,

low retention

VOYAGER - 50 μl – 12CH

A Multichannel reservoir – 10ml B FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl

C FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl D PCR cooling block 384_system

56

Conclusion

• In the context of a global pandemic where laboratories are under increasing pressure to

analyze more and more patient specimens to confirm COVID-19 cases, testing labs can

rapidly benefit from the advantages of the ASSIST PLUS pipetting robot, allowing them to

increase their sample processing capacity.

• Pipetting results were reproducible, precise and accurate, with no contamination

observed in controls or patient samples.

• The ASSIST PLUS pipetting robot, together with the VOYAGER adjustable tip spacing

pipette, increases sample processing capacity, reduces the need for manual intervention

by laboratory personnel and fast tracks assay development for COVID-19 testing.

• Low retention GRIPTIPS and a low dead volume SureFlo reagent reservoir allow the loss

of costly reagents, such as one-step RT-PCR master mix, to be reduced.

• The simple and fast ASSIST PLUS pipetting robot combined with the easy-to-use

VIALAB software, offers immediate help for testing labs.

• While the current protocol uses 384 well plates, it can be readily adapted to 96 well format

to meet future needs.

• Thanks to the VIALAB software, the pipetting programs can be easily adapted to any

specific protocols and labware.

CHAPTER 3: Application Notes

For more information

and a list of materials

used, please refer to

our website.

57

3.3 Increase your sample screening and genotyping

assay throughput with the VOYAGER adjustable

tip spacing pipette

Discover the advantages of setting up a genotyping assay

or sample screening with the VOYAGER adjustable

tip spacing pipette

Laboratories are continually facing the challenge of

increasing throughput in the most efficient and economical

way, to meet the need to process more and more samples

per day. Traditionally, handling and manipulating samples

between different labware formats involves the use of single

channel pipettes, especially in screening applications and

genotyping assays, which is slow, inefficient and error prone.

INTEGRA’s VOYAGER adjustable tip spacing pipette has enabled

scientists from the Technical University of Munich (TUM) to benefit

from the enhanced productivity of a multichannel pipette, reducing

tedious liquid handling tasks.

Compared to fully automated solutions, it provides seamless liquid

transfers between different standardized and non-standardized microplates,

tube and gel chamber formats, and can be used without any special training.

Tip spacing can be simply changed one-handedly with the push of a button,

eliminating the need for any manual adjustments.

The various operating modes of the VOYAGER adjustable tip spacing pipette help to speed

up monotonous pipetting steps, eliminate sample transfer errors between different labware

formats, and reduce the risk of developing repetitive strain injuries.

CHAPTER 3: Application Notes

58 CHAPTER 3: Application Notes

Key benefits

• The VOYAGER’s motorized adjustable tip

spacing enables the user to benefit from

the enhanced productivity of an electronic

multichannel pipette throughout the entire

genotyping assay, processing samples

faster than with traditional single channel

pipettes and helping to eliminate sample

transfer errors between different labware

formats.

• Tip spacing can be adjusted on the fly with

the push of a button to match different

types of labware, allowing the easy transfer

of multiple reaction mix samples from

microcentrifuge tubes directly to 96 or

384 well plates, and gel pockets.

• The availability of a range of pipetting

modes makes the VOYAGER a very

versatile and affordable tool to speed up

and standardize pipetting protocols.

• New users quickly get accustomed to the

electronic pipette thanks to its intuitive

design and easy-to-use pipetting modes.

Experimental set-up

In this protocol, two VOYAGER 8 channel adjustable tip spacing pipettes are used for a

genotyping set-up. The genotyping assay is based on a PCR method with a subsequent gel

electrophoresis.

The following protocol consists of sample transfers from 1.5 ml microcentrifuge tubes into a

96 well plate, and from a 96 well PCR plate into an agarose gel for electrophoresis.

Overview of the steps:

1. Template transfer

2. Sample transfer into the agarose gel

CHAPTER 3: Application Notes 59

Figure 1: Adjust the tip spacing by aligning it against the empty 96 well plate and tube rack.

Step-by-step procedure

1. Template transfer

Transfer the templates into a 96 well plate.

Use a VOYAGER 8 channel 300 μl electronic pipette

with 300 μl sterile, filter GRIPTIPS. Select ‘Tip spacing’

in the main menu of the pipette to set the required

spacing. Choose ‘Positions: 2’ in the tip spacing menu

and set the tip spacing according to the 96 well plate

and the microcentrifuge tubes in the rack (Figure 1).

Once saved, the tip spacing is available at any time, for

any other pipetting modes.

After saving the tip spacing, select ‘Pipet’ mode in the

main menu. Set your required sample transfer volume

and pipette the templates from the 1.5 ml microcentrifuge tubes into the 96 well plate (Figure

2). By pressing left and right on the Touch Wheel interface, the tip spacing can be adjusted on

the fly to fit each labware format.

Tips:

• Use the Repeat Dispense mode to dispense several samples successively if duplicate or

triplicate samples are required.

• Use the Pipet/Mix mode if samples require mixing in the target wells. Settings like mixing

cycles, pipetting speeds and volumes can quickly be adjusted.

Figure 2: Sample transfer from a microcentrifuge tube rack

to a 96 well plate.

60

2. Sample transfer into the agarose gel

Transfer the PCR product into the agarose gel.

After PCR, use the VOYAGER 8 channel 125 μl

electronic pipette with 125 μl sterile, filter GRIPTIPS to

transfer the samples from the 96 well PCR plate into the

agarose gel for subsequent gel electrophoresis (Figure

3). As in step 1, choose ‘Positions: 2’ in the tip spacing

menu and set the tip spacing according to the 96 well

PCR plate and the agarose gel.

Set the required sample volume as described in step 1

and transfer the samples from the PCR plate into the

agarose gel.

Tips:

• A low dispensing speed (e.g. 4) helps uniform filling of the wells in the agarose gel.

• If you want a controlled blowin – rather than automatic – keep the run button pressed while

dispensing. Blowin will occur when the run button is released.

CHAPTER 3: Application Notes

Figure 3: PCR product transfer into the agarose gel.

Conclusion

• The VOYAGER adjustable tip spacing pipette has enabled TUM researchers using

different labware formats to benefit greatly from the enhanced productivity of a

multichannel pipette, processing assays much faster than using a single channel pipette.

The tip spacing can be changed onehandedly at the touch of a button to fit different

labware formats, such as PCR plates, tubes and gel pockets.

• Thanks to the intuitive interface, users quickly become accustomed to the electronic

pipette. The different pipetting modes make the VOYAGER adjustable tip spacing pipette

a versatile yet affordable tool for working with labware of varying sizes and formats.

• The VOYAGER adjustable tip spacing pipette increases the speed of sample testing

set-ups, and helps eliminate sample transfer errors between different labware formats

and reduce the risk of developing repetitive strain injuries.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 3: Application Notes 61

3.4 PCR product purification with QIAquick® 96

PCR Purification Kit and the VIAFLO 96

handheld electronic pipette

Semi-automated PCR product purification on the

VIAFLO 96 handheld electronic pipette

QIAquick 96 PCR Purification Kit is suitable for purifying

up to 10 μg of material for downstream applications,

such as sequencing, cloning, labeling and microarrays.

The kit facilitates the removal of impurities like primers,

unincorporated nucleotides, buffers, salts, mineral oils,

agarose and enzymes. The vacuum-driven process is

much faster than centrifugation, and gives high,

reproducible yields. It is important to avoid

cross-contamination in nucleic acid purification,

and QIAGEN's column design is optimized to limit

carryover of contaminants. Although QIAquick 96

provides a high throughput solution, the elution,

washing and binding steps are very laborious and

time consuming if performed manually. With

VIAFLO 96 handheld electronic pipette, the hands-on time

is reduced, as samples and reagents can be transferred to

all 96 wells at once. This enables rapid and efficient, high throughput PCR clean-up.

Key benefits

• VIAFLO 96 and VIAFLO 384 allow

simultaneous pipetting of up to 96 or 384

wells, respectively, maximize throughput

of PCR purification by allowing transfer

samples and reagents in a single step.

• The z-heights can be predefined, choosing

the optimal value to prevent accidental

scratching of the well membrane for more

consistent results.

• Custom programming of the PCR product

clean-up steps allows pipetting parameters,

such as aspiration or dispensing speeds, to

be predefined. Prompt messages guide the

user through the entire pipetting protocol,

which is especially useful when several

pre-wetting steps are included.

• The VIAFLO 96 or VIAFLO 384's handsfree

automatic mode ensures that the

PCR clean up protocols are performed

in the same way each time, maximizing

reproducibility.

62 CHAPTER 3: Application Notes

Overview: How to purify PCR products with

VIAFLO 96

Experimental set-up

This protocol describes how PCR products

are purified using a VIAFLO 96 handheld

electronic pipette with a two position stage and

the QIAGEN QIAquick® 96 PCR Purification

Kit. The following procedure is based on the kit

manufacturer's protocol for purification of 96

samples (up to 10 μg PCR products).

A 96 channel pipetting head (50-1250 μl) is

used together with 1250 μl short, low retention,

sterile, filter GRIPTIPS. Customized VIALINK

programs are provided to perform the binding,

washing and elution steps. Before starting,

ethanol (96-100 %) should be added to the

Buffer PE concentrate.

Overview of the purification steps:

1. Step 1: Binding

2. Step 2: Washing

3. Step 3: Elution

The initial set-up of the QIAvac 96 Vacuum Manifold consists of a waste tray on top of a QIAvac

base, followed by a QIAquick 96 well plate (pink) mounted on a QIAvac 96 top plate, as shown in

Figure 1.

The QIAvac has to be attached to a vacuum source (house vacuum or vacuum pump) that

generates negative pressure between 100 and 600 mbar.

Figure 1: Initial set-up of the vacuum manifold.

CHAPTER 3: Application Notes 63

Step-by-step procedure

1. Binding

Binding the DNA to the silica-gel membrane.

Load the 1250 μl short, low retention, sterile, filter GRIPTIPS

on the VIAFLO 96. Place a 150 ml automation friendly reagent

reservoir in position A. The QIAvac 96 Vacuum Manifold

should be placed on position B of the VIAFLO 96 in landscape

orientation. No plateholder is needed on position B where the

manifold is placed.

Important: The vacuum manifold should be aligned before

each run (Figure 2).

Begin by launching the custom VIALINK program 'Qiaquick_

purification_M'. The pipette will prompt the user to place Buffer

PM on position A, then air is aspirated. This ensures that every

single drop of the liquid can be dispensed later. The

VIAFLO 96 will then guide the user through the two pre-wetting

steps, starting with aspiration and dispensing 200 μl of Buffer

PM. After a second aspiration, the pipette will display the prompt

'Move the head out of buffer', before dispensing the final 200 μl of

Buffer PM. This is followed by a 20 second wait, giving the buffer

residues time to flow down to the tip and be dispensed.

After pre-wetting, the pipette aspirates 75 μl Buffer

PM (three times the volume of the PCR product).

The instrument then tells the user to remove the

reservoir from position A, and replace it with the

96 well plate containing the 25 μl of PCR products.

After dispensing, and four mixing steps, the

resulting mixture is transferred to the QIAquick plate

wells in two steps. It is then time to switch on the

vacuum source, as indicated by the pipette.

Tips:

• Pre-wetting the tips prior to pipetting prevents

droplets and dripping when pipetting volatile

liquids, such as isopropanol, which is one of the

constituents in Buffer PM.

• Low retention GRIPTIPS (Figure 3) are used for these pipetting steps to avoid dripping.

Figure 2: Alignment of the QIAvac 96 Vacuum

Manifold.

Figure 3: Low retention versus standard tips.

64 CHAPTER 3: Application Notes

2. Washing

Two-step purification of the PCR product.

Eject the used tips and load new 1250 μl short, low retention, sterile, filter GRIPTIPS on the

VIAFLO 96. Place a new 300 ml automation friendly reagent reservoir in position A. The

VIAFLO 96 will then prompt the user to pour Buffer PE into the reservoir, followed by a prewetting

step, which is necessary since the buffer contains ethanol. After pre-wetting, the pipette

will aspirate 900 μl of Buffer PE, and dispense it into QIAquick plate wells. The instrument will

then notify the user that is it time to turn on the vacuum pump. With the pump turned on, another

dose of the buffer is dispensed into the wells, followed by a 10 minute wait to dry the membrane

and remove all residual ethanol.

Important: The final drying step is crucial to remove residual ethanol prior to elution.

Residual ethanol in the elution buffer could inhibit downstream applications (e.g. PCR).

Tip: After this step, the manufacturer suggests tapping the plate on a stack of absorbent paper

to ensure that all residual buffer is removed.

3. Elution

Elution of DNA from the silica-gel membrane.

When prompted, start by replacing the waste tray with the

blue collection microtube rack provided, which contains

1.2 ml vessels (Figure 4a). Load new 1250 μl short, low

retention, sterile, filter GRIPTIPS, and place a new

150 ml automation friendly reagent reservoir in position A.

The instrument will then prompt the user to place Buffer

EB into the reservoir, aspirate 80 μl, and dispense it into

the QIAquick plate wells. After a 1 minute incubation, the

pipette tells the user to switch on the vacuum source for

5 minutes.

Tips:

• The purified PCR product could also be eluted in

a 96 well microplate. In this case, when replacing the

waste tray, the 96 well microplate has to be placed on

the empty blue collection tube rack (Figure 4b).

• For increased DNA concentration, decrease the elution

volume to 60 μl, as per QIAGEN's recommendations, in

the VIALINK software.

Figure 4: Elution into a) provided collection

microtubes or b) a 96 well microplate.

A)

B)

CHAPTER 3: Application Notes 65

Remarks

Vacuum manifold:

Alignment of the vacuum manifold is very important in this process. Adding marks on the deck

helps to reposition the manifold whenever needed. To check the position of the well plate on top

of the vacuum manifold, attach the tips manually to the pipette. The pipette tips should be in the

middle of the wells. If not, adjust the position of the vacuum manifold on the deck.

Automatic mode:

The VIAFLO 96 can also operate in hands-free automatic mode, allowing the user to have

more walk-away time and less interaction, which is highly beneficial when using the instrument

in a laminar flow cabinet. The customized automatic VIALINK program can be found on the

INTEGRA website.

Conclusion

• The VIAFLO 96 electronic handheld pipette allows fast and simple liquid transfers for high

throughput PCR product purification.

• Optimized pipette settings enable accurate sample and reagent transfer, without the tip

touching and scratching the QIAquick membrane.

• The VIAFLO 96 electronic handheld pipette's compact design takes up minimal space

and fits on any lab bench.

• The unique operating concept makes the VIAFLO 96 and VIAFLO 384 as easy to use as

a conventional electronic pipette.

• The QIAvac 96 manifold is easily placed on the instrument and allows the processing of

other kits using 96 well silica-membrane or filter plates.

• Another option for this application is the MINI 96, which is the most affordable 96 channel

option on the market.

For more information

and a list of materials

used, please refer to

our website.

66 CHAPTER 3: Application Notes

3.5 PCR purification with Beckman Coulter

AMPure XP magnetic beads and the VIAFLO 96

Automatic magnetic bead purification with the VIAFLO 96

handheld electronic pipette

Agencourt AMPure XP magnetic beads (Beckman Coulter) are an efficient

way to clean up samples for PCR, NGS, cloning and microarrays. The kit

provides a solution for medium to high throughput requirements when carried

out in a 96 well plate, but the protocol involves many washing and transfer

steps that make it tedious to perform manually. With the VIAFLO 96,

a handheld 96 channel electronic pipette, multistep protocols such as

PCR clean-up and DNA purification can be

performed quickly and efficiently, increasing

throughput tremendously by transferring

samples and reagents to all 96 wells at once.

Thanks to its unique operating concept,

the VIAFLO 96 remains as easy to use as

a traditional handheld pipette and can even

provide critical information (user-defined

prompts) about the protocol steps.

Key benefits

• The VIAFLO 96 enables transfer of

samples, reagents and wash solutions to

96 wells at once, increasing the throughput

of magnetic bead-based DNA purification

methods.

• The partial tip loading of the VIAFLO 96

allows purification of fewer than 96 DNA

samples if necessary; 8, 16, 24, 32, 40

or 48 GRIPTIPS can be loaded for easy

purification of different numbers of samples.

• The optimal immersion depth for removing

supernatant or adding liquid right onto

the samples is guaranteed by defining the

z-height of the VIAFLO 96.

• The Tip Align setting of the VIAFLO 96

automatically positions the tips in the center

of the wells of a 96 well plate, avoiding any

disturbance of the beads.

CHAPTER 3: Application Notes 67

Overview: How to automate PCR purification steps

with VIAFLO 96

The VIAFLO 96 handheld electronic pipette with a three position stage is used to purify DNA

with AMPure XP beads from Beckman Coulter. The following protocol is an example of a set-up

for 96 samples, where each well of a 96 well plate is filled with 10 μl of DNA sample and 18 μl

of AMPure XP beads, then further processed with the VIAFLO 96. The PCR purification can

be performed manually or semi-automated using the VIAFLO 96 in automatic mode.

Custom-made VIALINK programs are provided. The VIALINK programs are set up according

to the manufacturer’s protocol (AMPure XP Beckman Coulter).

Step-by-step procedure

1. Dispense AMPure XP beads into PCR tubes

Transfer AMPure XP beads from the stock solution into 12 PCR tubes placed in

a cooling block from INTEGRA.

Note: The cooling block is just used as a support in this instance, not for cooling down the

samples.

To ensure a homogenous stock solution, beads are thoroughly mixed by shaking/inverting until

the solution appears consistent in color. The beads are transferred into 12 PCR tubes using

the Repeat Dispense mode of a VIAFLO single channel 1250 μl electronic pipette. A customized

VIALINK program (AMP_Transfer1) is available to aid bead transfer.

For optimal pipetting, ensure beads are thoroughly mixed before each transfer. Mixing steps

can be defined by the number of cycles and the pipetting speed. Both influence the efficiency

of mixing and thus the quality of the

clean-up. Saving these parameters in the

pipetting program ensures that mixing is

always carried out as defined, yielding

consistent results. Insert a pre- and

post-dispense step to enhance accuracy

and precision while pipetting precious

reagents, such as AMPure XP beads.

Tip: The use of sterile, filter, low retention

GRIPTIPS ensures that every dispense

is as accurate as possible, with no loss of

beads or sample.

Figure 1: Transfer AMPure XP beads from the stock solution into 12 PCR

tubes.

68 CHAPTER 3: Application Notes

2. Transfer AMPure XP beads into the DNA samples

Transfer AMPure XP beads from the PCR tubes into a 96 well plate preloaded

with DNA samples.

Pipette the beads from the PCR tubes

into the 96 well plate using a VIAFLO

12 channel 50 μl electronic pipette. For

optimal pipetting, make sure the tips are

exchanged, and mix the beads thoroughly

before each transfer. A customized

VIALINK program (AMP_Transfer2) is

provided for this step.

Tip: Use low retention GRIPTIPS to

minimize loss of beads adhering to the

tip wall.

3. Mixing and binding of the AMPure XP beads

Mixing and binding of the magnetic beads to the PCR samples.

Load GRIPTIPS (position A) then select

and run the AMPure_XP_M program on

the VIAFLO 96. The samples are now

mixed 10 times by pipetting up and down

on position B. A five minute wait time

follows, timed by the VIAFLO 96, to allow

the DNA to bind to the beads.

Tip: Use the z-height setting of the

VIAFLO 96 to define the optimal tip

immersion depth. This prevents air

entering the tip during mixing and avoids

the pipette tip touching the bottom of the

plate. Setting the Tip Align support strength to 3 for positions A and B makes it more comfortable

to use the VIAFLO 96. These settings can be incorporated into the program so that they are not

forgotten.

Figure 2: Transfer AMPure XP beads from the PCR tubes into a 96 well

plate preloaded with DNA samples.

Figure 3: Mixing and binding of the magnetic beads to the PCR samples.

CHAPTER 3: Application Notes 69

4. Magnetic separation of the AMPure XP beads

Separating the magnetic beads from the PCR samples.

Note: Make sure new GRIPTIPS are loaded

before continuing the protocol to ensure

removal of the supernatant without bead

carryover.

A prompt on the pipette screen reminds the

user to move the sample plate from position

AB onto the 96 well magnet (position B) and

place an automation friendly reagent reservoir

for waste collection on position AB. After a two

minute incubation time, the beads form a ringshaped

structure and the solution becomes

clear. Load new GRIPTIPS before continuing the procedure to ensure accurate removal of

the supernatant without bead carryover. Follow the instructions on the pipette and aspirate

the supernatant slowly from the sample, dispensing it into the waste reagent reservoir

(position AB).

Tip: To avoid disturbing the ring of beads, the supernatant is aspirated slowly at speed 1.

Leave 5 μl of supernatant in the plate to prevent beads being drawn out during aspiration.

The z-height limit is again used to ensure that the beads are not disturbed during pipetting.

5. AMPure XP bead clean-up

Wash the magnetic beads twice with 70 % ethanol.

Place an automation friendly reagent reservoir

containing 70 % ethanol on position A and

change the GRIPTIPS before continuing

with the wash step. Follow the prompts on

the pipette. Pre-wet the GRIPTIPS with 70 %

ethanol. Then wash the samples with 70 %

ethanol. Repeat the washing step again as

indicated by the pipette.

Tip: Pre-wetting the GRIPTIPS with 70 %

ethanol ensures equilibration of the humidity

and the temperature between the air in the

pipette/tips and the sample/liquid. In-house testing has shown that low retention GRIPTIPS

prevent ethanol from dripping while traveling from one pipetting position to another.

Figure 4: Separating the magnetic beads from the PCR samples.

Figure 5: Wash the magnetic beads twice with 70 % ethanol.

70

6. Elute samples from the magnetic beads

Elute the purified samples from the magnetic beads by adding the elution

buffer.

As indicated by the pipette, replace the 70 %

ethanol reagent reservoir on position A with

an elution buffer reagent reservoir and move

the sample plate from the magnet (position B)

to position AB. Load new GRIPTIPS before

continuing with the protocol. After transferring

and thoroughly mixing the elution buffer with

the beads, the pipette prompts the user to

place the sample plate back onto the magnet

(position B). During the one minute incubation

time, place a new 96 well plate on position AB.

7. Transfer the sample eluates

Transfer the sample eluates into the new 96 well plate.

Note: Load new GRIPTIPS to ensure a clean

eluate transfer without bead carryover.

Continue with the same program, slowly and

carefully transferring the eluates from position

B into the new plate (position AB).

Tip: Optimizing pipette settings (aspiration

speed, volume and height) allows the volume

of the transferred eluate to be maximized

without carryover of beads. These settings

can be easily tweaked at any time. Performing

a test run with water before implementing any

new assay is an ideal way to optimize pipette settings.

Figure 6: Elute samples from the magnetic beads.

Figure 7: Transfer the sample eluates into the new 96 well plate.

CHAPTER 3: Application Notes

CHAPTER 3: Application Notes 71

Remarks

Automatic mode:

The VIAFLO 96 can also operate on its own,

enabling less user interaction, which in turn

improves ergonomics and reproducibility. This

also makes it even more ideal for use in tight

spaces, such as under a laminar flow cabinet.

Partial tip load:

If you are not working with a full set of 96

samples, the VIAFLO 96 is able to work with

any number of tips loaded, allowing purification

of smaller numbers of samples. Figure 8: Automatic mode and partial tip load.

Conclusion

• The VIAFLO 96 is perfectly suited to magnetic bead purification in a 96 well format. An

entire plate with 96 samples can be purified in a fraction of the time it would take with a

traditional pipette.

• Optimized tip immersion and pipette settings in combination with the use of low retention

GRIPTIPS allow maximum sample recovery at the end of the purification protocol.

• The VIAFLO 96 can guide the user through the entire protocol step by step, ensuring the

correct workflow and enhancing the reproducibility of results.

• The optional automatic mode of the VIAFLO 96 enables the instrument to operate on its

own to minimize pipetting errors, making it even more ideal for use under a laminar flow

cabinet.

For more information

and a list of materials

used, please refer to

our website.

72 CHAPTER 3: Application Notes

3.6 PCR purification with Beckman Coulter

AMPure XP magnetic beads and

the ASSIST PLUS

Automatic magnetic bead purification with

ASSIST PLUS pipetting robot

Agencourt AMPure XP beads (Beckman Coulter) are used

for DNA purification in a variety of applications, including PCR,

NGS, cloning and microarrays. The ASSIST PLUS pipetting

robot provides a solution for optimal bead

separation and maximized recovery of

precious samples. User guidance

throughout the entire protocol

ensures an error-free pipetting

procedure. Careful and accurate

handling of the magnetic beads

by the ASSIST PLUS leads to

superior reproducibility and consistency

during the experiment. Taken together, the

ASSIST PLUS provides researchers with an easy

and highly efficient way to purify DNA from PCR reactions using AMPure XP magnetic beads.

Key benefits

• The VIAFLO and VOYAGER electronic

pipettes, in combination with

ASSIST PLUS, provide unmatched

pipetting ergonomics.

• Optimal pipette settings, including tip

immersion depth, pipetting speeds and

angles, maximize reproducibility and

sample recovery.

• Exact positioning of the pipette tips in the

sample wells avoids the risk of disturbing

the ring of magnetic beads or bead

carryover.

• The ASSIST PLUS automates many steps

of a magnetic bead purification protocol

and guides the user through the remaining

manual operations to ensure an error-free

process.

CHAPTER 3: Application Notes 73

Overview: How to automate PCR purification steps

with ASSIST PLUS

The ASSIST PLUS is used to purify DNA samples using AMPure XP beads (Beckman Coulter).

The pipetting robot runs a VOYAGER 8 channel 125 μl electronic pipette with 125 μl sterile,

filter, low retention GRIPTIPS. The use of low retention GRIPTIPS guarantees optimal liquid

handling of viscous (AMPure XP buffer) and volatile (70 % ethanol) solutions.

Below is an example set-up for 24 samples, preparing 10 μl DNA samples (position B) with

18 μl of AMPure XP beads (position A). The pipetting programs were prepared according to the

manufacturer’s protocol (AMPure XP, Beckman Coulter) using VIALAB software.

The protocol is divided into two programs that guide the user through every step of the PCR

purification process.

• Program 1: Binding (AMP_BINDING)

• Program 2: Washing and elution (AMP_WASH_ELUTE)

Experimental set-up: Program 1

Deck position A: PCR 8 tube strip containing the AMPure XP

beads (Figure 1, blue), placed onto a cooling

block from INTEGRA. Note: the cooling block

is just used as a support in this instance, and

not for cooling down the samples.

Deck position B: 96 well plate with 24 DNA samples for

purification (Figure 1, green).

Deck position C: 96 well ring magnet.

74 CHAPTER 3: Application Notes

Figure 1: Pipetting schema, set-up for program 1.

A B C

Run program 1: transfer & binding

Select and run the AMP_BINDING program on the VOYAGER electronic pipette. The

ASSIST PLUS pipetting robot immediately starts the protocol.

1. AMPure XP transfer

Transferring AMPure XP beads from an 8 tube PCR strip to a 96 well plate

containing the DNA samples.

To ensure the AMPure XP buffer is homogenous, the beads are resuspended by pipetting up

and down 10 times before being transferred to the samples. The beads and DNA fragments

are thoroughly mixed together before the pipette automatically starts the timer for a 5 minute

incubation, ensuring optimal conditions for the DNA strands to bind onto the magnetic beads.

Tip: Using low retention GRIPTIPS rather than regular GRIPTIPS prevents the loss of AMPure

XP beads during the pipetting steps (see Figure 2).

VOYAGER 8 channel

125 μl

50/125 μl sterile, filter,

low retention GRIPTIPS

PCR 8-Tube Strip on cooling

plate – 200 μl

96 well plate Sapphire

– 200 μl

96 well plate Sapphire on 96 well ring

magnet – 200 μl

CHAPTER 3: Application Notes 75

Figure 2: The image highlights the advantages of using low retention GRIPTIPS versus regular

GRIPTIPS when pipetting AMPure XP beads.

Figure 3: The beads and DNA fragments are thoroughly mixed together before the incubation.

76 CHAPTER 3: Application Notes

2. Magnetic separation of the AMPure XP beads

Separating the magnetic beads from the PCR samples.

A message instructs the user to move the plate (position B) onto the magnet (position C).

Continue the program to start the timer. After a two minute incubation on the magnet the

beads form a ring in the sample well and the solution becomes clear. The program resumes

automatically, and the supernatant is removed. On completion of this step, the pipette prompts

the user to continue with the AMP_WASH_ELUTE program and to replace the labware on

position A with the 8 row polypropylene (PP) reagent reservoir containing the ethanol and

elution buffer.

Tip: The supernatant is aspirated slowly using the Tip Travel feature of the ASSIST PLUS

to avoid disturbing the ring of beads. The Tip Travel feature keeps the tip immersion depth

constant during aspiration and dispensing. 5 μl of supernatant remain in the plate to prevent

beads being drawn out during aspiration.

Figure 4: The ASSIST PLUS settings allow removal of the supernatant without any bead carryover.

CHAPTER 3: Application Notes 77

Experimental set-up: Program 2

Deck position A: The 96 well PCR cooling block is replaced by

an 8 row polypropylene (PP) reagent reservoir

filled with 70 % ethanol in row 1 (blue) and

elution buffer in row 2 (orange). Row 8 is used

for waste (purple).

Deck position B: Emtpy 96 well plate.

Deck position C: 96 well ring magnet and 96 well plate with 24

DNA samples for purification (green).

Figure 5: Pipetting schema, set-up for program 2.

VOYAGER 8 channel

125 μl

50/125 μl sterile, filter,

low retention GRIPTIPS 8 row reagent reservoir 96 well plate Sapphire

– 200 μl

96 well plate Sapphire on 96 well ring

magnet – 200 μl

A B C

78 CHAPTER 3: Application Notes

Run program 2: Washing & elution

Start the AMP_WASH_ELUTE program on the VOYAGER electronic pipette. The

ASSIST PLUS washes the beads twice by automatically adding and removing ethanol.

3. Magnetic bead clean-up

Washing the magnetic beads twice with 70 % ethanol.

The programmed pipette settings allow the beads to be washed without disturbing the bead

ring. At the end of the second washing step, all the ethanol is removed. If necessary, an

additional drying time can easily be added using VIALAB software.

Tip: The use of low retention GRIPTIPS prevents ethanol from dripping while traveling from

position A to position C (see Figure 6).

Figure 6: The image highlights the advantages of using low retention GRIPTIPS (left) versus regular

GRIPTIPS (right) when pipetting ethanol.

4. Elute samples from the magnetic beads

Eluting the samples from the magnetic beads by adding an elution buffer.

The pipette prompts the user to move the reaction plate from the magnet (position C) to position

B. Continuing the protocol, the ASSIST PLUS transfers the elution buffer to the DNA samples

bound to the magnetic beads (position B, orange). After mixing carefully and thoroughly 10

times, the pipette prompts the user to place the 96 well plate on the magnet (position C).

CHAPTER 3: Application Notes 79

5. Transfer the sample eluates

Transferring the sample eluates into a new 96 well plate.

As indicated by the pipette, place a new 96 well plate onto position B and continue the program.

The sample eluates are then transferred into the new plate automatically.

Tip: Optimized pipette settings (aspiration speed, volume, height, tip travel and tip touch) allow

the volume of eluate transferred to be maximized without carryover of beads (see Figure 6).

A tip touch after the transfer removes droplets that may still cling to the end of the pipette tips.

Pipetting heights on the ASSIST PLUS can be fine-tuned at any time. Performing a test run with

water before implementing any new assay is an ideal way to optimize pipette settings.

Results

Figure 7: Magnetic beads are clearly visible in the 96 well plate with no supernatant remaining.

80 CHAPTER 3: Application Notes

Figure 8: No carryover of beads is observed in the eluate.

Conclusion

• Magnetic bead purifications can be easily automated on the ASSIST PLUS pipetting

robot.

• Optimized tip immersion and pipette settings together with the use of low retention

GRIPTIPS allow maximum sample recovery at the end of the purification protocol.

• The pipette loaded onto the ASSIST PLUS prompts the user when needed, eliminating

the risk of human errors.

• VIALAB programs can be easily adapted to specific labware.

• Prolonged pipetting tasks lead to repetitive strain injury. This can be avoided by

automating these steps with the ASSIST PLUS.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 4: Customer Testimonials 81

CHAPTER 4:

Customer testimonials

Our range of innovative liquid handling products has helped countless laboratories to achieve

PCR success, improve their throughput and further their ground-breaking research. But don’t

just take our word for it! Here are a few stories from our satisfied customers, demonstrating why

INTEGRA Biosciences is the right choice for PCR pipetting solutions and labware.

4.1 INTEGRA pipettes – the obvious choice for

start-up PCR labs

The gradual reopening of the world following the pandemic has led to an unprecedented

demand for COVID-19 testing, with schools, universities and workplaces relying on negative

PCR tests to continue operating. Matrix Diagnostics – a dedicated COVID-19 testing lab in

California – is helping to fulfill this critical need, relying on INTEGRA’s EVOLVE and MINI 96

pipettes to streamline and accelerate PCR workflows.

PCR-based diagnostic testing is a well-established technique in clinical labs around the world,

and this method has been brought to the attention of every household as the gold standard for

COVID-19 testing. However, the public is less aware that the sensitivity of this technique makes

it time-consuming and troublesome to perform without the right tools, as it is very sensitive to

pipetting errors and cross-contamination.

Founded in January 2021, Matrix Diagnostics

was established to meet the growing demand

for PCR testing in the San Francisco Bay Area,

and the newly formed team understood the need

for effective pipetting solutions from the outset.

Fady Ettnas, Lab Manager at Matrix Diagnostics,

explained: “We realized that, to meet the

anticipated demand for testing, we would have to

turnover between 2000 and 5000 samples every

day. This seemed like an impossible task for a new

lab with limited resources but, after implementing

INTEGRA’s pipettes in our lab, we quickly

alleviated the pipetting bottlenecks, putting us on

track to achieve our targets.”

Photo courtesy of Matrix Diagnostics

82

Evolving workflows

“Our protocols involve a range of repetitive

pipetting steps – including mixing reagents

and serial dilutions – for thousands of

samples a day, which has the potential

to be a cumbersome and error-prone

task,” Fady continued. “We therefore

chose INTEGRA’s EVOLVE manual

pipettes and MINI 96 portable electronic

pipettes to improve the reproducibility

and productivity of our workflows. We

have a number of single channel EVOLVE

pipettes, covering volumes ranging from

0.2 to 5000 μl, as well as 8, 12, and 16

channel models. What I like most about

EVOLVE is its ergonomic design and

ability to set volumes in a flash. The

unique design of INTEGRA’s GRIPTIPS also means that they never leak or fall off, avoiding

cross-contamination and maintaining sterility. We also use the compact MINI 96 extensively,

which is especially well suited to PCR set-up. It saves a lot of time and effort – around 15

minutes per cycle – when performing the wash steps. And because we run more than 25 cycles

every day, this is a huge saving, allowing us to process a much higher number of samples. It is

a perfect and affordable solution for our needs.”

A long-term investment

The benefits of these pipettes to users, particularly in terms of preventing physical strain

caused by repeated pipetting actions, are a priceless advantage. “I think the pipettes are a

great investment with huge returns, allowing the team to process more samples and improving

their pipetting experience. The company’s customer service is quick, responsive and helpful

and, crucially, the team was able to advise us on the right choice of pipettes to meet our

workload and objectives.”

Planning future with INTEGRA

“Currently, we are only offering COVID-19 tests, but we plan to expand to include other tests

including sexually transmitted diseases, urinary tract infections and flu, and we know that we

will need to automate our workflow. We will need something flexible and incredibly efficient and,

therefore, we are planning to acquire an ASSIST PLUS pipetting robot. I like all the INTEGRA

products that I’ve used, and have rarely encountered even minor technical issues. I think they

are the most obvious pipetting choice for both for start-ups and established lab set-ups, and are

well worth the investment,” Fady concluded.

Photo courtesy of Matrix Diagnostics

CHAPTER 4: Customer Testimonials

83

Photo courtesy of Harvard Medical School

4.2 A better qPCR pipetting experience

Manual pipetting can be a major bottleneck for research laboratories, especially when they face

the challenge of combining accurate results with high throughput. Like all repetitive tasks that

require precise actions, filling multiwell plates by hand is time consuming, and physically and

mentally draining, which can lead to errors. When Daisy Shu joined the Saint-Geniez laboratory

at Harvard Medical School, her experience was quite different, thanks to the INTEGRA VIAFLO

electronic pipettes.

From patients to pipettes

After graduating in optometry from the University of New South

Wales in Sydney, Daisy worked as an optometrist for two years

before deciding to pursue a PhD in cataract research at the

University of Sydney. She explained: “The move from my usual

clinical work with patients to research was a big change for me,

as I had to dive deep into molecular biology. I didn't even know

how to use a pipette back then! Cataracts – clouding of the

eye’s lens – are a leading cause of blindness worldwide, and I

studied their formation and ways to prevent that happening. My

focus was on transforming growth factor beta (TGF-β), which

has an important role in cancer metastasis, but is also relevant

for certain types of cataracts. I looked at the different signaling

pathways it activates and how those pathways interlink.”

Daisy completed her PhD in January 2019, and straight

afterwards flew to Boston to work as postdoctoral fellow in the

Saint-Geniez laboratory, continuing her research into eye health.

Here, she was able to apply her knowledge of TGF-β to agerelated

macular degeneration (AMD). Daisy continued: “I'm now

looking at how TGF-β causes the retinal mitochondria to change morphology and become

dysfunctional, altering cellular metabolism. The research is still at an early stage, so we're

mainly trying to understand how to prevent AMD, but the end goal is to find a cure.”

A better pipetting experience

At Harvard, Daisy was introduced to VIAFLO electronic pipettes, which were a complete

contrast to the large, fully automated pipetting workstation she had used during her PhD

research. The laboratory was already using two VIAFLO pipettes – a 125 μl eight channel

pipette and a 12.5 μl single channel version – and their flexibility compared to the automated

workstation dramatically improved her pipetting experience. “Complete automation on a large

workstation has its place, but there are downsides,” said Daisy. “You have to program every

CHAPTER 4: Customer Testimonials

84

single step perfectly before you can click one button and run the

protocol, and the process of fine-tuning takes a long time.”

“I found the VIAFLO pipettes amazing. A lot of our work is PCRbased,

performed in 384 well plates, and the VIAFLO pipettes

are real lifesavers. I use the 8 channel VIAFLO for most qPCR

liquid transfers, and the single channel pipette to add the

primers. Once you've made your master mixes and programmed

the pipette, it's really fast; it only takes me 20 minutes to

do a complete 384 well plate. When I was using the robotic

workstation in Sydney, I used to think that doing a qPCR was

really a big deal. Now, with the INTEGRA pipettes, it's just

so easy.”

VIAFLO pipettes provide a choice of pipetting modes and allow

easy adjustment of parameters such as volume and speed, as

well as providing pre-set programs and the option for custom

workflows. This helps laboratories to reduce errors and increase

throughput and reproducibility regardless of the users’ pipetting

experience. For Daisy, VIAFLO electronic pipettes have become the standard for how pipetting

should be: “In any pipetting workflow, you have to get every step right first time, otherwise you’d

end up having to troubleshoot the assay and do it again. I'm really surprised when I hear people

from other labs say they pipette each well individually with manual single channel pipettes. I’m

sure that would take forever compared to electronic pipetting, and my eyes would really suffer.

The VIAFLOs make everything easy. I love the color coding – it makes it so simple to match the

right tip to the right pipette – and the instrument can even be set to alert you when you need to

pipette again.”

CHAPTER 4: Customer Testimonials

Photo courtesy of Harvard Medical School

85

4.3 COVID-19 – Accelerate your PCR set-up

The emergence and outbreak of the novel coronavirus SARS-CoV-2 (COVID-19) has placed

unprecedented demands on laboratories testing patient samples for COVID-19, leaving

scientific staff to contend with a spiraling influx of COVID-19 samples and a rapid, continuous

growth in workload. Among the challenges faced by the Microbiology and Molecular Pathology

Department at Sullivan Nicolaides Pathology (SNP) – part of the Sonic Healthcare Group

– in Brisbane, Australia, is the increased pressure on laboratory automation used for both

coronavirus and pre-existing respiratory virus panel testing.

As a result of the coronavirus pandemic, SNP found itself analyzing extreme numbers of

samples, which exhausted the capacity of its automation platforms. At the same time, staff

were faced with a need to spend more time working up new virus testing protocols, which

were often performed manually or using semi-automated methods to fast track test response

times, leaving them prone to increased ergonomic strain. There was a clear need for additional

automated liquid handling instruments to increase sample processing capacity, reduce manual

intervention by laboratory analysts and fast track assay development for COVID-19 sample

testing.

Working together

In early March 2020, Kelly Magin and James Sundholm from

INTEGRA’s Australian distributor, BioTools Pty Ltd, partnered

with Shane Byrne, Scientific Department Head, Microbiology and

Molecular Pathology Department, SNP, to support COVID-19 testing

of patient samples using the ASSIST PLUS pipetting robot. An

ASSIST PLUS automated pipetting protocol was developed and

validated, enabling samples to be prepared in low volume, 384 well

plates for subsequent processing on a rapid, high throughput,

plate-based, real-time PCR amplification and detection instrument.

A VOYAGER adjustable tip spacing pipette and low retention

GRIPTIPS were used to transfer one-step RT-PCR master mix from a

low dead volume (<20 μl) SureFlo 10 ml reagent reservoir into a 384

well plate. The VOYAGER pipette also allowed automatic transfer

and reformatting of nucleic acid template extracted from combined

nasopharyngeal/oropharyngeal flocked swab(s) or sputum samples,

from 4 x FluidX™ 1.0 ml 96 format tube racks into the 384 well plate. The total PCR reaction

volume was reduced to 10 μl; 7.5 μl one-step RT-PCR master mix and 2.5 μl of nucleic acid

template. This miniaturization doubled the available testing capacity and simultaneously

reduced consumption of expensive one-step RT-PCR reagents of dwindling availability, with

associated cost savings.

Photo courtesy of Sullivan Nicolaides

Pathology

CHAPTER 4: Customer Testimonials

86

Defining success

SNP successfully validated the automated

protocol against its existing manual

processing method, performed using a

handheld electronic pipette. The results

were shown to be reproducible, precise

and accurate, with no contamination

observed in either the control or patient

samples. The compact, easy-to-use

ASSIST PLUS pipetting robot, complete

with validated protocol, was fully deployed

within five working days. While the current

protocol uses 384 well plates, it can be

readily adapted to 96 well format to meet

future needs.

4.4 Reducing protocol time for PCR using

96 channel pipette

Implementing an INTEGRA VIAFLO 96 electronic pipette has enabled the Virus- and Prion

Validation (VPV) Department at Octapharma Biopharmaceuticals GmbH, (Frankfurt, Germany)

to reduce the time taken to undertake PCR assays by greater than 60 %.

Since its foundation in 1983, Octapharma has been committed to patient care and medical

innovation. Its core business is the development and production of human proteins from human

plasma and human cell-lines.

The VPV Department has been set-up to investigate pathogen inactivation and removal steps

along the manufacturing processes. Among other techniques, multi-step 96 well format PCR

assays were developed, which involve three washing steps twice in the protocol. To undertake

their PCR assay more efficiently, Octapharma sought a system that enabled reproducible and

accurate liquid handling in the 96 well format and was able to completely remove residual liquid

as well as avoid well-to-well contamination.

Dr. Andreas Volk, a research scientist at Octapharma Biopharmaceuticals commented: "The

classical liquid handling solutions, fully automated robots or ELISA plate washers were either

too costly or prone to cross contamination in a PCR assay." He added: "When we tested the

INTEGRA VIAFLO 96 channel pipette, it fully met our requirements as it enabled mediumthroughput

liquid handling while minimizing cross-contamination. Additionally, the

Photo courtesy of Sullivan Nicolaides Pathology

CHAPTER 4: Customer Testimonials

87

VIAFLO 96 electronic pipette provided all the

adjustment options, which we had been used

to with manual pipettes, plus a specified tip

immersion depth for each pipetting step. With

our PCR protocol, which involves ten full liquid

transfers per plate, we now only use half the

amount of pipette tips as we can use the same

tips for liquid addition and aspiration in each

washing step. VPV Department staff has found

using the VIAFLO 96 benchtop pipette highly

intuitive and the overall time required for our

PCR washing procedures has been reduced to

approximately one third of the original time."

The INTEGRA VIAFLO 96 is a handheld 96

channel electronic pipette that has struck a

chord with scientists looking for fast, precise

and easy simultaneous transfer of

96 samples from microplates without the

cost of a fully automated system. The

VIAFLO 96 was designed to be handled just

like a standard handheld pipette – a fact

borne out by consistent end user feedback

that no special skills or training are required to

operate it. Users immediately benefit from the

increased productivity delivered by their VIAFLO 96. Fast replication or reformatting of 96 and

384 well plates and high precision transferring of reagents, compounds and solutions to or from

microplates with the VIAFLO 96 is as easy as pipetting with a standard electronic pipette into

a single tube. Four pipetting heads with pipetting volumes up to 12.5 μl, 125 μl, 300 μl or

1250 μl are available for the VIAFLO 96. These pipetting heads are interchangeable within

seconds enabling optimal matching of the available volume range to the application performed.

For 384 well pipetting, an enhanced version is available with VIAFLO 384. It features

384 channel pipetting heads in the volume range of 12.5 μl and 125 μl and is compatible with

96 channel pipetting heads.

Dr. Andreas Volk, Octapharma Biopharmaceuticals

CHAPTER 4: Customer Testimonials

88

CHAPTER 5:

Conclusion

So, there you have it, a full run down of PCR. By now, you should have all the information you

need to become a PCR pro, but if you’d still like to learn more about this interesting topic, we

have a wealth of articles on our website. Whatever your PCR requirements, we at INTEGRA

Biosciences are always available to answer your questions and provide you with the best

workflow solutions.

CHAPTER 5: Conclusion

89

CHAPTER 6:

References

1.1 The complete guide to PCR

1. Crow, E. (2012). Mind Your P's And Q's: A Short Primer On Proofreading Polymerases.

https://bitesizebio.com/8080/mind-your-ps-and-qs-a-short-primer-on-proofreadingpolymerases

2. Kim, S. W. et al. (2008). Crystal structure of Pfu, the high fidelity DNA polymerase from

Pyrococcus furiosus. International Journal of Biological Macromolecules, 42(4), 356-

361. https://doi.org/10.1016/j.ijbiomac.2008.01.010

3. ThermoFisher Scientific (n.d.). PCR Setup – Six Critical Components to Consider.

https://www.thermofisher.com/ch/en/home/life-science/cloning/cloning-learningcenter/

invitrogen-school-of-molecular-biology/pcr-education/pcr-reagents-enzymes/

pcr-component-considerations.html

4. AAT Bioquest (2020). What is the function of MgCl2 in PCR?

https://www.aatbio.com/resources/faq-frequently-asked-questions/What-is-thefunction-

of-MgCl2-in-PCR

5. Lorenz, T. C. (2012). Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting

and Optimization Strategies. Journal of Visualized Experiments, 63, e3998.

https://doi.org/10.3791/3998

6. Merck (n.d.). Polyermase Chain Reaction.

https://www.sigmaaldrich.com/CH/en/technical-documents/technical-article/

genomics/pcr/polymerase-chain-reaction

7. Viana, R. V., Wallis, C. L. (2011). Good Clinical Laboratory Practice (GCLP) for

Molecular Based Tests Used in Diagnostic Laboratories. In Akyar, I. (Ed.), Wide

Spectra of Quality Control (29-52). InTech.

https://cdn.intechopen.com/pdfs/23728/InTech-Good_clinical_laboratory_

practice_%20gclp_for_molecular_based_tests_used_in_diagnostic_laboratories.pdf

8. Ogene M. (2021). How does ddPCR work?

https://mogene.com/how-does-ddpcr-work

9. ThermoFisher Scientific (2016). Real-time PCR handbook.

https://www.ffclrp.usp.br/divulgacao/emu/real_time/manuais/Apostila%20qPCRHandbook.

pdf

CHAPTER 6: References

90

10. Prediger, E. (2017). Digital PCR (dPCR) – What is it and why use it?

https://eu.idtdna.com/pages/technology/qpcr-and-pcr/digital-pcr

11. Bio-Rad Laboratories (n.d.). Introduction to Digital PCR.

https://www.bio-rad.com/en-uk/life-science/learning-center/introduction-to-digital-pcr

12. Bio-Rad Laboratories (n.d.). Digital PCR and Real-Time PCR (qPCR) Choices for

Different Applications.

https://www.bio-rad.com/en-uk/life-science/learning-center/digital-pcr-and-real-timepcr-

qpcr-choices-for-different-applications

13. Schoenbrunner, N. J. et al. (2017). Covalent modification of primers improves PCR

amplification specificity and yield. Biology Methods and Protocols, 2(1).

https://doi.org/10.1016/j.ijbiomac.2008.01.010

14. Merck (n.d.). Hot Start PCR.

https://www.sigmaaldrich.com/CH/en/technical-documents/technical-article/

genomics/pcr/hot-start-pcr

15. Parichha, A. (2021). Nested PCR || Principle and usage.

https://www.youtube.com/watch?v=nHCjgo2Ze0o

16. New England Biolabs (n.d.). FAQ: What is touchdown PCR?

https://international.neb.com/faqs/0001/01/01/what-is-touchdown-pcr

17. Parichha, A. (2021). Touch down PCR.

https://www.youtube.com/watch?v=s9oV2-53esA

18. Cheriyedath, S. (2018). History of Polymerase Chain Reaction (PCR).

https://www.news-medical.net/life-sciences/History-of-Polymerase-Chain-Reaction-

(PCR).aspx

19. Arney, K. (2020). The Story of PCR.

https://geneticsunzipped.com/news/2020/11/3/the-story-of-pcr

20. Biosearch Technologies (2022). Taq facts.

https://blog.biosearchtech.com/thebiosearchtechblog/bid/48174/taq-facts

21. National Museum of American History (n.d.). Mr. Cycle, Thermal Cycler.

https://americanhistory.si.edu/collections/search/object/nmah_1000862

CHAPTER 6: References

91

1.2 Simple PCR tips that can make or break your success

1. Cheriyedath, S. (2018). History of Polymerase Chain Reaction (PCR).

https://www.news-medical.net/life-sciences/History-of-Polymerase-Chain-Reaction-

(PCR).aspx

2. Seeding Labs (2019). How To: PCR Calculations.

https://www.youtube.com/watch?v=CnQV5_CEvAo

3. McCauley, B. (2020). Setting Up PCR Reactions.

https://brianmccauley.net/bio-6b/6b-lab/polymerase-chain-reaction/pcr-setup

4. New England Biolabs (n.d.). Guidelines for PCR Optimization with Taq DNA

Polymerase.

https://international.neb.com/tools-and-resources/usage-guidelines/guidelines-forpcr-

optimization-with-taq-dna-polymerase

5. Lorenz, T. C. (2012). Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting

and Optimization Strategies. Journal of Visualized Experiments, 63, e3998.

https://doi.org/10.3791/3998

6. Gold Biotechnology (2020). How To: PCR Master Mixes.

https://www.youtube.com/watch?v=LSfvCJ9gUQU

1.3 Setting up a PCR lab from scratch

1. Bustin, S. A., Benes, V., Garson, J. A., et al (2009). The MIQE Guidelines: Minimum

Information for Publication of Quantitative Real-Time PCR Experiments. Clinical

Chemistry, 55(4), 611–622.

https://doi.org/10.1373/clinchem.2008.112797

2. National Human Genome Research Institute (2020). Polymerase Chain Reaction

(PCR) Fact Sheet.

https://www.genome.gov/about-genomics/fact-sheets/Polymerase-Chain-Reaction-

Fact-Sheet

3. Viana, R. V., Wallis, C. L. (2011). Good Clinical Laboratory Practice (GCLP) for

Molecular Based Tests Used in Diagnostic Laboratories.

https://cdn.intechopen.com/pdfs/23728/InTech-Good_clinical_laboratory_practice_

gclp_for_molecular_based_tests_used_in_diagnostic_laboratories.pdf

4. Redig, J. (2014). The Devil is in the Details: How to Setup a PCR Laboratory.

https://bitesizebio.com/19880/the-devil-is-in-the-details-how-to-setup-a-pcrlaboratory

5. Mifflin, T. E. (n. d.). Setting Up a PCR Laboratory.

https://pubmed.ncbi.nlm.nih.gov/21357132/

CHAPTER 6: References

92

6. Gu, M. (n. d.). Molecular Laboratory Design And Its Contamination Safeguards.

https://www.scimmit.com/molecular-laboratory-design-and-its-contaminationsafeguards

7. Lee, R. (2015). Molecular Laboratory Design, QA/QC Considerations.

https://www.aphl.org/programs/newborn_screening/Documents/2015_Molecular-

Workshop/Molecular-Laboratory-Design-QAQC-Considerations.pdf

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work

1. Bustin, S. A., Benes, V., Garson, J. A. et al. (2009). The MIQE guidelines: minimum

information for publication of quantitative real-time PCR experiments. Clinical

Chemistry, 55(4), 611-622.

https://doi.org/10.1373/clinchem.2008.112797

2. Rutledge, R. G., Côté, C. (2003). Mathematics of quantitative kinetic PCR and the

application of standard curves. Nucleic Acids Research, 31(16).

https://www.gene-quantification.de/rudledge-2003.pdf

3. Applied biological materials (2016). Polymerase chain reaction (PCR) – Quantitative

PCR (qPCR).

https://www.youtube.com/watch?v=YhXj5Yy4ksQ

4. Nagy, A., Vitásková, E., Černíková, L. et al. (2017). Evaluation of TaqMan qPCR

system integrating two identically labelled hydrolysis probes in single assay. Scientific

reports, 7.

https://doi.org/10.1038/srep41392

5. Bradburn, S. (n.d.). How to calculate PCR primer efficiencies.

https://toptipbio.com/calculate-primer-efficiencies

6. Bio-Rad (n.d.). qPCR assay design and optimization.

https://www.bio-rad.com/en-ch/applications-technologies/qpcr-assay-designoptimization?

ID=LUSO7RIVK

7. University of Western Australia (2016). Melt curve analysis in qPCR experiments.

https://www.youtube.com/watch?v=FvJnXKzejSQ

8. Bio-Rad (2011). Real time QPCR data analysis tutorial.

https://www.youtube.com/watch?v=GQOnX1-SUrI

9. Bio-Rad (2011). Real time QPCR data analysis tutorial (part 2).

https://www.youtube.com/watch?v=tgp4bbnj-ng

10. Kannan, S. (2021). 4 easy steps to analyze your qPCR data using double delta Ct

analysis.

https://bitesizebio.com/24894/4-easy-steps-to-analyze-your-qpcr-data-using-doubledelta-

ct-analysis

CHAPTER 6: References

93

1.5 How to design primers for PCR

1. Benchling (n.d.). Primer Design.

https://www.benchling.com/primers

2. Addgene (n.d.). How to Design a Primer.

https://www.addgene.org/protocols/primer-design

3. PREMIER Biosoft (n.d.). PCR Primer Design Guidelines.

http://www.premierbiosoft.com/tech_notes/PCR_Primer_Design.html

4. Merck (n.d.). Oligonucleotide Melting Temperature.

https://www.sigmaaldrich.com/CH/en/technical-documents/protocol/genomics/pcr/

oligos-melting-temp

5. Integrated DNA Technologies (n.d.). How do you calculate the annealing temperature

for PCR?

https://eu.idtdna.com/pages/support/faqs/how-do-you-calculate-the-annealingtemperature-

for-pcr

CHAPTER 6: References

INTEGRA Biosciences AG

7205 Zizers, Switzerland

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info@integra-biosciences.com

INTEGRA Biosciences Nordic ApS

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www.integra-biosciences.com

HOW TO BECOME A PCR PRO

The polymerase chain reaction (PCR) is a key life sciences technique. It has been used in

molecular biology – including molecular diagnostics – for many years, and a number of different

types, for example, RT-PCR, qPCR, vPCR and ddPCR have been developed over time.

Today, PCR is a vital tool for the detection of pathogens, such as the SARS-CoV-2 virus, and

is essential for genotyping and NGS library preparation. However, PCR is well known for being

difficult to run successfully and several parameters must be considered when planning the PCR

protocol.

We have therefore compiled this eBook – consisting of in-depth educational articles, relevant

app notes and customer testimonials – to help you understand how PCR works, and what needs

to be considered to perform effective PCR reactions. We also demonstrate how our solutions

can help you to enhance the throughput of your lab, and become a PCR pro in no time.

Dr Éva Mészáros

Application Specialist

eva.meszaros@integra-biosciences.com

Anina Werner

Content Manager

anina.werner@integra-biosciences.com

FOREWORD

TABLE OF CONTENTS

CHAPTER 1: What you need to know about PCR

1.1 The complete guide to PCR 2

1.2 Simple PCR tips that can make or break your success 15

1.3 Setting up a PCR lab from scratch 20

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work 24

1.5 How to design primers for PCR 32

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 38

CHAPTER 3: Application notes

3.1 Efficient and automated 384 well qPCR set-up with the ASSIST PLUS pipetting robot 42

3.2 Automated RT-PCR set-up for COVID-19 testing 49

3.3 Increase your sample screening and genotyping assay throughput with the VOYAGER 57

adjustable tip spacing pipette

3.4 PCR product purification with QIAquick® 96 PCR Purification Kit and the 61

VIAFLO 96 handheld electronic pipette

3.5 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 66

VIAFLO 96

3.6 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 72

ASSIST PLUS

CHAPTER 4: Customer testimonials

4.1 INTEGRA pipettes – the obvious choice for start-up PCR labs 81

4.2 A better qPCR pipetting experience 83

4.3 COVID-19 – Accelerate your PCR set-up 85

4.4 Reducing protocol time for PCR using 96 channel pipette 86

CHAPTER 5: Conclusion 88

CHAPTER 6: References 89

2

CHAPTER 1:

What you need to know about PCR

In this chapter, we will cover topics such as PCR’s fascinating history, its mechanism and

different variations, and techniques for troubleshooting common issues you may encounter.

We’ll also go through tips for establishing a PCR lab, as well as a comprehensive overview of all

things related to qPCR and primer design.

1.1 The complete guide to PCR

Polymerase chain reaction (PCR) methods have been carried out in labs around the world since

the 1980s, opening the door for an array of new applications, such as genetic engineering,

genotyping and sequencing. In this article, we take a deep dive into this fascinating technique

by explaining its mechanism, exploring its history, looking into the different types of PCR,

discussing troubleshooting tips and much more.

CHAPTER 1: What you need to know about PCR

3

What is PCR?

The polymerase chain reaction (PCR) is a fast and inexpensive technique for amplifying a DNA

sequence of interest. It consists of three steps:

• Denaturation: The sample is heated to separate the DNA into two single strands.

• Annealing: The temperature is lowered to allow primers to anneal to specific single-stranded

DNA segments, flanking the sequence to be amplified.

• Extension: The temperature is raised to the optimum working temperature of the polymerase

enzyme, which then makes a complementary copy of the DNA sequence of interest.

One such repetition or 'thermal cycle' theoretically doubles the amount of the DNA sequence of

interest present in the reaction. Typically, 25 to 40 cycles are performed – resulting in millions

or even billions of DNA copies – depending largely on the amount of DNA in the starting sample

and the number of amplicon copies needed for post-PCR applications.

The three steps of a PCR reaction are carried out automatically by a thermal cycler, but can

only be successful if the master mix has been correctly prepared. The following sections

explain the components that make up the master mix and how they interact with the template

DNA during thermal cycling.

PCR master mix components

The PCR master mix consists of six components:

• PCR-grade water: Certified to be free of contaminants, nucleases and inhibitors.

• dNTPs: Containing equal concentrations of the four nucleotides (dATP, dCTP, dGTP and

dTTP), which are the 'building blocks' to create complementary copies of the DNA sequence

of interest.

• Forward and reverse primers: Short, single-stranded DNA sequences that anneal

specifically to the plus and minus strands of the template DNA, flanking the sequence to

be amplified. For some assays – such as protocols amplifying much-studied genes or DNA

sequences of common bacteria – ready-to-use primers can be purchased. However, many

experiments require custom PCR primers tailored to the region of interest of the template DNA

and the reaction conditions.

CHAPTER 1: What you need to know about PCR

4

• DNA polymerase: Taq-polymerase is the most commonly used enzyme for PCR reactions.

It uses dNTPs to create complementary copies of the DNA sequence of interest. For some

applications, such as mutagenesis, Taq-polymerase is not accurate enough and the use

of high fidelity polymerases is recommended. Just like Taq-polymerase, they sometimes

add an incorrect nucleotide when replicating the template DNA but, as they have a 3' to

5' exonuclease activity, they 'proofread' the newly synthesized strands and correct any

mistakes.1 This proofreading step is highly beneficial for accuracy but it also slows down PCR

reactions, and high fidelity polymerases (also called slow polymerases) therefore need about

twice the time of Taq-polymerase to create a complementary DNA strand. The most popular

high fidelity DNA polymerase is Pfu-polymerase.2

• Buffer: Provides a suitable environment for the DNA polymerase, with a pH between 8.0

and 9.5.3

• Magnesium chloride: Increases the activity of the DNA polymerase and helps primers

to anneal to the template DNA for a higher amplification rate.4 This cofactor is sometimes

included in the buffer in a sufficient concentration.5

The template DNA , which may be genomic DNA (gDNA), complementary DNA (cDNA) or

plasmid DNA (pDNA), is then added after master mix preparation.

The 3 steps of PCR

After preparing the PCR master mix and adding the template DNA samples to it, you can load

your reaction tubes, PCR strips or microplates into the thermal cycler. They will then go through

the following steps:

• Denaturation: The thermal cycler first heats the reaction mix to 95-98 °C to denature the

template DNA, separating it into two single strands. Depending on your sample, this usually

takes 2-5 minutes during the first thermal cycle, and 10-60 seconds for subsequent cycles.

• Annealing: When the temperature is lowered, the primers anneal to the sequences flanking

the template DNA region of interest. Depending on the sequence and melting temperature of

your primers, this step usually takes 30-60 seconds, and the optimal annealing temperature

typically lies between 45 and 60 °C.

• Extension: The temperature is increased to 72 °C, which is the ideal working temperature

for the Taq-polymerase. Depending on the synthesis rate of your polymerase, and the length

of the target sequence, it usually takes 20-60 seconds to create complementary copies

of the DNA sequence of interest.6 After approximately 25-40 cycles – depending on the

amount of DNA present at the start, and the number of amplicon copies needed for post-PCR

applications7 – the last extension step should be extended to 5 minutes or longer, allowing the

Taq-polymerase to finish the synthesis of uncompleted amplicons.5 If you can't immediately

take your samples out of the thermal cycler after the final extension step because you're busy

with other experiments, program it to cool your samples to 4 °C. For overnight runs where you

CHAPTER 1: What you need to know about PCR

5

leave your samples in the thermal cycler for hours after the final extension step, you should

opt for a holding temperature of 10 °C instead of 4 °C, as it causes less wear and tear on your

machine.

As shown in the image above, the amount of PCR product theoretically doubles at every

thermal cycle, leading to an exponential increase of PCR product. However, in reality, the phase

of exponential amplification eventually levels off and reaches a plateau because the reagents

have been consumed and the DNA polymerase activity decreases.

The different types of PCR

After performing a standard PCR reaction, you can determine the concentration, yield and

purity of the amplified DNA sequences using gel electrophoresis, spectrophotometry or

fluorometry. However, you can’t determine the amount of template DNA present in a sample

before amplification using standard PCR. If this is a requirement for your experiment, you have

to perform a qPCR reaction.

qPCR

qPCR – also called real-time PCR, quantitative PCR or quantitative real-time PCR – is a

technique used to detect and measure the amplification of target DNA as it is produced.

In contrast to conventional PCR reactions, qPCR requires a fluorescent intercalating dye

or fluorescently-labeled probes, and a thermal cycler that can measure fluorescence and

calculate the cycle threshold (Ct) value. Typically, the fluorescence intensity increases

proportionately to the concentration of the PCR product being formed, measuring quantities

of the target in real time.

CHAPTER 1: What you need to know about PCR

6

qPCR can be divided into dye-based methods (e.g. SYBR® Green) and probe-based methods

(e.g. TaqMan®).

RT-PCR and RT-qPCR

Another limitation of standard PCR is that it can only be used to amplify DNA sequences. If you

want to amplify RNA target sequences, you have to use RT-PCR.

RT-PCR

vPCR

Reverse transcription PCR (RT-PCR) is used to amplify RNA target sequences, such as

messenger RNA or RNA virus genomes. This type of PCR involves an initial incubation of

the RNA samples with a reverse transcriptase enzyme and a DNA primer – comprising

sequence-specific oligo (dT)s or random hexamers – prior to the PCR amplification.

For viability PCR (vPCR), each sample needs to be split into two aliquots. One aliquot is

incubated with a photoreactive intercalating dye that is unable to diffuse through intact cell

membranes of live cells. This means that it only intercalates into the DNA of dead cells. When

this aliquot is subsequently treated with a blue light, the dye binds irreversibly to the DNA. Both

aliquots are then subject to DNA purification and qPCR amplification. If they exhibit similar

qPCR signals, the target microorganisms in the sample are mostly viable. If the dye-treated

aliquot exhibits a weaker signal, the target microorganisms are mostly dead. vPCR is an

important technique in diagnostics, agriculture and food safety.

You can also perform a qPCR reaction instead of executing a standard PCR reaction after

the reverse transcription step, which produces cDNA from RNA. This PCR variant is called

RT-qPCR.

vPCR

The third limitation of standard PCR is that it cannot distinguish between the DNA of viable

and non-viable cells. You should use vPCR if this is important to your application, for example,

because you want to know if the infectious microorganisms in a clinical sample are dead or

alive.

CHAPTER 1: What you need to know about PCR

7

ddPCR

Digital droplet PCR (ddPCR) is another relatively new type of PCR. It uses fluorescently labeled

probes to detect DNA sequences of interest, and a water-oil emulsion system to split each

sample into about 20,000 nanoliter-sized droplets. After amplification, every droplet of the

sample is analyzed on its own. Droplets that contain at least one DNA sequence of interest emit

a fluorescent signal – and are consequently positive – while droplets without the DNA sequence

of interest don't fluoresce, and are therefore negative. Using the Poisson distribution, you can

then determine the concentration of the DNA sequence of interest in the original sample by

analyzing the ratio of positive to negative droplets for absolute quantification.8

An advantage of ddPCR compared to qPCR is that it's more precise. While qPCR can detect

two-fold differences in DNA target sequence variation, e.g. discriminate 1 copy from 2 copies

of a gene, ddPCR can discriminate 7 copies from 8 copies, which means that it can detect

differences as small as 1.2-fold.9 On top of that, ddPCR is better suited for multiplexing assays

if you want to determine the ratio of low abundance to high abundance DNA sequences of

interest, such as rare mutations on wild type backgrounds. When using qPCR, the fluorescent

signal from the high abundance sequences can dominate and obscure the signal from the

low abundance sequences. This risk is ruled out with ddPCR, as each droplet behaves as

an individual PCR reaction and contains either zero, one or, at most, a few sequences of

interest.10,11

ddPCR

CHAPTER 1: What you need to know about PCR

8

Due to these advantages, ddPCR is often preferred over qPCR for the detection of mutations

and SNPs (single nucleotide polymorphisms), allelic discrimination, gene expression studies,

and the analysis of copy number variations.12

Hot start PCR

If your PCR reaction results in non-specific amplification, you can try to increase the reaction

specificity using a hot start polymerase. This enzyme remains inactive during master mix

preparation and sample addition at room temperature, eliminating the risk that unintended

products and primer dimers are formed during PCR set-up.13

Nested and semi-nested PCR

Nested or semi-nested PCR are alternatives to hot start PCR that increase reaction specificity.

Nested PCR uses two sets of primers and two successive PCR reactions. The first set of

primers is designed to amplify a DNA sequence slightly longer than the sequence of interest.

During the second PCR reaction, the second set of primers that is specific to the sequence of

interest anneals to the amplicons of the first PCR reaction and helps to amplify the sequence of

interest.14,15

Nested PCR

CHAPTER 1: What you need to know about PCR

9

Semi Nested PCR

Semi-nested PCR works similarly to nested PCR. During the first PCR reaction, one primer

anneals to the sequence of interest and the second primer to a region flanking the sequence of

interest. This primer is then replaced with a second primer annealing to the region of interest

during the second PCR reaction.

The idea behind nested and semi-nested PCR is that, if non-specific products were amplified

during the first PCR reaction, these will not be amplified during the second PCR reaction, as the

primers cannot anneal to them.

Touchdown PCR

A third type of PCR developed to increase reaction specificity is touchdown PCR. The assay

set-up for touchdown PCR is identical to the set-up for standard PCR. The only difference lies in

the annealing step. During the first thermal cycle, the annealing temperature should be several

degrees above the optimal primer annealing temperature, then be lowered by 1-2 °C every

second cycle.16 These high temperatures during the first cycles avoid PCR primers forming

primer-dimers or binding to regions outside the DNA sequence of interest. The downside is

that the PCR primers don't all sufficiently bind to the template DNA, which leads to low yields.17

However, this can be tolerated, as the low yield of specific amplicons is then exponentially

amplified with every thermal cycle that is performed at the optimal annealing temperature.

CHAPTER 1: What you need to know about PCR

10

The history of PCR

As we've shown, there are many different types of PCR, and some of them have only recently

been developed. However, the foundation for PCR was laid in the 1950s:

• In 1953, James Watson and Francis Crick discovered the double-helix structure of DNA, and

suggested that there might be a possible copying mechanism for DNA.

• Four years later, Arthur Kornberg identified the first DNA polymerase that was able to copy the

template DNA, although only in one direction.

• In 1971, Gobind Khorana and his team started to work on DNA repair synthesis. Their

technique used DNA polymerase repeatedly, but employed only a single primer template

complex, which did not allow exponential amplification.

• At the same time, Kjell Kleppe from Khorana's lab proposed a two primer system that would

double the amount of DNA in a sample, but no one actually conducted the experiment to

find out whether it worked. The reason for this was probably that there was not yet a DNA

polymerase that could withstand the high temperatures of the denaturation step. This means

that they would have had to add a fresh dose of enzyme after every thermal cycle.

• In 1983, Kary Mullis, working at Cetus Corporation, added a second primer to the opposite

strand, and realized that repeated use of DNA polymerase triggers a chain reaction that will

amplify a specific DNA sequence, thus inventing PCR. The patent got approved in 1987, and

he won the Nobel Prize in Chemistry six years later.

• In 1976, the thermostable enzyme Taq-polymerase – which is typically used in PCR today

– was first isolated from the bacterium Thermus aquaticus, which had been discovered in a

hot spring of Yellowstone National Park in 1969. When it was finally incorporated into PCR

workflows in 1988, it removed the need to add a new dose of enzyme after every thermal

cycle, paving the way for the invention of automated thermal cyclers.18,19,20

CHAPTER 1: What you need to know about PCR

11

PCR troubleshooting

One of the most important troubleshooting mechanisms is to always include positive and

negative control samples.

If the sequence of interest wasn't amplified in your positive control sample, your master mix,

template DNA or thermal cycler could be the source of the problem:

• Master mix: Have you added the right volume and concentration of each reagent, and have

you cooled your reagents during master mix preparation?

• Template DNA: Have you run an agarose gel to ensure that your template DNA isn't

degraded? Is your template DNA pure enough and, if not, have you purified it?

• Thermal cycler: Is the number of thermal cycles sufficient for your assay? Have you

programmed the device correctly, and is it calibrated to ensure that it performs the reaction

steps at the right temperatures?

If the sequence of interest was amplified in your negative control sample, one or more

components of your master mix is contaminated. PCR reactions are very sensitive, and create

large number of copies of DNA sequences from minute amounts of starting material, so

contamination is a common issue. To prevent it, the right lab set-up is crucial.

CHAPTER 1: What you need to know about PCR

12

Lab set-up

Ideally, your PCR lab should have two rooms, each divided into two areas. The first room should

be exclusively used for pre-PCR activities, and divided into a master mix preparation area and a

sample preparation area. The second room should have a dedicated area for amplification, and

another one for product analysis.

If you’re lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible. In addition to the spatial separation, you could also consider setting up your PCR

reactions in the morning, and performing the amplification and analysis steps in the afternoon.

Temporally separating the different steps of your PCR reactions may limit your flexibility and

make you lose some time, but lowers the risk of aerosols with high DNA concentrations from the

analysis area contaminating your master mix and samples in the pre-PCR area.

On top of these precautionary measures, you should always work in biosafety cabinets or

laminar flow hoods when setting up your PCR reactions, use different sets of pipettes for master

mix preparation, sample preparation and analysis, and make sure that you use filter tips and

consumables that are free of DNase, RNase and PCR inhibitors.

Specificity

Another major PCR challenge is specificity. As explained before, it can be improved using hot

start, nested, semi-nested or touchdown PCR. A further option to prevent the amplification of

regions outside the DNA sequence of interest, as well as the formation of secondary structures,

is to redesign your primers.

CHAPTER 1: What you need to know about PCR

13

Use this checklist to see whether your primers meet all the requirements:

• Are your primers between 18 and 24 bp long?

• Is your target sequence length between 100 and 3000 bp for standard PCR assays, or 75

and 150 bp for qPCR assays?

• Do your primers have melting temperatures between 50 and 60 °C, and within 5 °C of

each other?

• Have you performed a gradient PCR to determine the optimal annealing temperature?

• Does the GC content of your primers lie between 40 and 60 %?

• Have you avoided runs or repeats of four or more bases or dinucleotides?

• Have you made sure that your primers are not homologous to a template DNA sequence

outside the region of interest?

• Have you checked that your primers can't form stable secondary structures?

PCR equipment

The most important PCR instrument is certainly the thermal cycler but, as the right pipetting

devices can help create faster and more efficient workflows with fewer errors, we'll also look at

a few different liquid handling options in this section.

Thermal cyclers

Before the development of thermal cyclers, scientists had to manually move their samples

between water baths of different temperatures. The first thermal cycler prototype called 'Mr.

Cycle' also used water baths to heat and cool the samples, and was developed by engineers

at Cetus Corporation, where Kary Mullis worked when he invented PCR.21 Today's instruments

work with electric heating and refrigeration units, and many different models with various

additional features are available.

For standard PCR, a thermal cycler that can heat and cool your samples to the required

temperatures might be sufficient to complete the different reaction steps. However, your

thermal cycler will need additional properties – such as gradient capability or an integrated

fluorometer – if you want to perform gradient PCR assays to optimize primer annealing

temperatures, or qPCR assays to determine the amount of template DNA present in a sample

before amplification.

CHAPTER 1: What you need to know about PCR

14

Pipettes

While the thermal cycler is the star of PCR labs, the right pipettes help you to process more

samples in less time, while ensuring maximal accuracy and precision. Electronic pipettes

offering a Repeat Dispense mode, for example, are a great option to boost the efficiency of

aliquoting master mix into an entire well plate. Adjustable tip spacing pipettes, paired with low

dead volume reagent reservoirs, can be a useful alternative to single channel pipettes when

transferring reagents and samples between different labware formats. And, if you want to

significantly cut your PCR set-up and purification time, pipetting robots or 96 and 384 channel

pipettes might be the right tool for you.

Conclusion

We hope that this article has been useful in helping you understand the mechanisms behind

the different types of PCR, and has shown you different ways to avoid contamination and nonspecific

amplification.

CHAPTER 1: What you need to know about PCR

15

1.2

Since the outbreak of the COVID-19 pandemic, PCR is on everybody's lips. However, only

people working in the lab know how difficult it can be to get the desired results using this wellestablished

technique. Out of this frustration came the popular joke that PCR should stand for

’pipette, cry, repeat’. To ensure that this stays a joke from now on, and that your PCR reactions

never drive you to despair again, we have compiled the most important tips and tricks for a

successful PCR set-up.

What is PCR?

The polymerase chain reaction (PCR) is used to amplify specific DNA sequences for

downstream use. Its inventor Kary Mullis, whose patent on PCR was approved in 1987, was

awarded the Nobel Prize in Chemistry six years later,1 and since this time, PCR has remained

one of the most essential molecular biology techniques. Genetic engineering, genotyping,

sequencing and the identification of familial relationships, to name a few examples, wouldn't be

possible without it.

PCR tips and tricks

To perform PCR reactions, you need to prepare a master mix, add template DNA, and amplify

the sequence of interest using a thermal cycler. This might seem straightforward, but it is far

from it. Calculating the required amounts of master mix reagents correctly to get the right

volume, at the right concentration, is the first challenge.

Once this is accomplished, the reagents need to be mixed together. The difficulty here is that

the liquids usually have to be cooled and they are often highly viscous, sticky and needed

in minimal quantities. In addition, work must be performed in a concentrated manner, as

distractions or interruptions can quickly lead to a situation where you no longer know which

reagents have already been added to the master mix. Errors such as skipping a tube or well can

CHAPTER 1: What you need to know about PCR

Simple PCR tips that can make or break your success

16

also easily occur when filling PCR strips or plates with master mix and adding template DNA,

especially when using single channel pipettes.

The last and probably biggest challenge is to keep your PCR reactions free from contamination.

PCR is a very sensitive assay that can create a large number of nucleic acid copies from a tiny

amount of starting material, so amplicon and sample contamination can be a huge problem.

Master mix calculations

Let's first have a look at the mathematical calculations needed to set up a PCR master mix.

We'll assume that you want to set up several PCR reactions with a volume of 50 μl each.

To calculate the required volume for each reagent, it is best to create a table (see Table 1) with

the necessary components, and fill in the stock concentrations and desired final concentrations

for the buffer, the MgCl2, the dNTPs and the primers. Then, calculate the dilution factors by

dividing the stock concentration by the final concentration. To determine the volume needed for

a single PCR reaction, divide the desired reaction volume by the dilution factor.2

For the polymerase, a slightly different equation is needed. The manufacturer of the enzyme

will tell you the amount of polymerase in one μl, e.g. 5 Units/μl. Fill in this value in the column

for the stock concentration and put the desired amount – e.g. 1.25 Units – in the column for

the final concentration. The volume needed can then be calculated as follows: 1.25 Units x

(1 μl / 5 Units) = 0.25 μl.3

The template DNA volume required depends on your sample type. You should add about 1 pg

to 10 ng of plasmid or viral DNA, and 1 ng to 1 μg of genomic DNA. In the example below, we

calculated how much you would need to use for 0.5 μl of a 1 μg/μl template DNA.4

Finally, add the required volumes for all the reagents. The difference between the desired total

reaction volume (50 μl) and the result obtained gives you the volume of PCR-grade water.5

REAGENT STOCK CONC. FINAL CONC.

(CF)

DILUTION

FACTOR

(= STOCK

CON. / CF)

VOLUME NEEDED

(= 50 ΜL / DIL.

FACTOR)

Buffer 10X 1X 10 5 μl

MgCl2 25 mM 1.5 mM 16.66 3 μl

dNTPs 10 mM 0.2 mM 50 1 μl

Forward primer 10 μM 250 nM 40 1.25 μl

Reverse primer 10 μM 250 nM 40 1.25 μl

Polymerase 5 Units/μl 1.25 Units - 0.25 μl

Template DNA 1 μg/μl - - 0.5 μl

PCR-grade water - - - 37.75 μl

Table 1: Example of a PCR master mix table

CHAPTER 1: What you need to know about PCR

17

After determining the required reagent volumes for one PCR reaction, you can simply multiply

them by your sample number (plus the negative and positive controls) to get the total volumes

for the entire PCR set-up. We recommend adding one additional aliquot to that result, as some

of the master mix may be lost during pipetting due to evaporation, adherence to the tip, or

pipetting inaccuracies.

That's it, you are now ready to set up your PCR reactions by following the best pipetting

practices listed below.

Best PCR pipetting practices

Start by preparing your master mix from all the components listed above, except the template

DNA. The huge advantage of preparing the entire quantity of master mix needed for an

experiment, and subsequently transferring single aliquots into PCR strips or plates, is that

you can pipette higher volumes with better accuracy. On top of that, it reduces pipetting steps,

making the entire process less tiring and error prone. Since pipetting mistakes cannot be

completely ruled out, you should add the master mix components in order of their price, starting

with the most affordable reagent. This way, you waste less money if you have to start over.6

Once your master mix is finished, well mixed and dispensed into tubes or plates, you can

add the template DNA. As the DNA samples are usually highly viscous and needed in small

quantities, you should either dispense them into the master mix or onto the wall of the tube or

well. After dispensing, keep the plunger depressed while dragging the tip gently along the wall

of your labware to remove any residual liquid. In addition, we recommend using low retention

tips.

If you're not using a hot start polymerase, cool your reagents throughout the entire process of

master mix preparation and sample addition, to prevent non-specific amplification.

When you are ready to load your samples into the thermal cycler, check that they are tightly

capped or sealed, and spin them down to ensure that no droplets remain on the labware wall

during amplification.

Pipetting solutions for PCR reactions

Before discussing various pipetting solutions, we would like to address one of the most

important aspects of liquid handling. No matter which pipettes you choose, ensure that they are

well maintained by regularly calibrating them and checking their performance in between uses.

The most affordable pipettes for master mix preparation would be manual single channel

models. However, as you need to accurately measure and mix several very expensive

reagents, we recommend investing in electronic single channel pipettes. The motor-controlled

piston movement guarantees that they always dispense the exact desired volume, minimizing

variability to increase the precision and accuracy of pipetting.

For the container, you can either prepare the master mix in a tube or, if you intend to transfer

it with an electronic multichannel pipette, in a low dead volume reagent reservoir. The

CHAPTER 1: What you need to know about PCR

18

ASSIST PLUS pipetting robot transferring master mix into a 384 well PCR plate

combination of an electronic multichannel pipette and a reservoir is ideal for this step, because

you can fill several tubes or wells simultaneously. On top of that, electronic multichannel

pipettes usually feature a Repeat Dispense mode, allowing you to aspirate a large volume

of master mix, then dispense it into multiple smaller aliquots. It is also possible to use an

electronic single channel pipette if you have a low throughput.

To add template DNA to the master mix aliquots, an adjustable tip spacing pipette can be

very handy if the labware format of your samples doesn't match the container used for PCR

amplification. For example, it allows you to transfer several template DNA samples from

microcentrifuge tubes to an entire row or column of a 96 well plate in one step.

High throughput labs might even want to take advantage of automated solutions for master

mix plating and sample transfer, such as a pipetting platform that is capable of automating

electronic pipettes.

CHAPTER 1: What you need to know about PCR

19

How to prevent PCR contamination

Several preventative measures should be taken to avoid contaminating your master mix or

template DNA with amplicons that were generated during previous PCRs.

One of the most effective means is to physically separate the master mix preparation, template

DNA addition, amplification and analysis areas from one another, and to work in laminar flow

or biosafety cabinets. Each work zone, and its corresponding equipment, should be cleaned

before and after an experiment, and tools used in one area should never enter another one.

When it comes to consumables, make sure you purchase sterile products that are certified to

be free from DNase, RNase and PCR inhibitors. Pipette tips should form a perfect seal with

the pipette to eliminate contamination that may occur when tips drip or fall off. Using filter tips

will also avoid the risk of aerosols entering your pipettes and contaminating subsequent PCR

reactions.

As you're a potential source of contamination too, always wear gloves to prevent introducing

enzymes, microbes and skin cells to the reaction, and change them when going from one area

to another. On top of that, keep your tubes closed whenever possible during the entire PCR

set-up.

Despite these preventative measures, you can't completely eliminate the possibility of

contaminated PCR reactions. To avoid having to throw away your entire stock of a certain

reagent if this occurs, prepare single use aliquots of your master mix components. You can also

prepare aliquots of positive and negative controls, as well as serial dilutions of standards for

quantitative PCR (qPCR) assays, ahead of time. Electronic pipettes with repeat dispense and

serial dilute modes can be helpful for this task, not only to reduce the risk of contamination, but

also to increase the efficiency of PCR set-up.

Conclusion

PCR is a fundamental technique in research, diagnostics and forensics. It often involves

pipetting minuscule reagent volumes with tricky properties, so it can be difficult to obtain the

desired results. On top of that, contamination can have a huge impact on results, as it's a very

sensitive assay. We hope that the tips and tricks provided in this article will help you make

your future PCR reactions a success. Many of these recommendations can also be applied to

other amplification assays, such as reverse transcription and qPCR, loop-mediated isothermal

amplification (LAMP) and helicase-dependent amplification (HDA).

CHAPTER 1: What you need to know about PCR

20

1.3 Setting up a PCR lab from scratch

PCR reactions are very sensitive and create a large number of copies of nucleic acids

from minute amounts of starting material. This makes them a fundamental and highly

effective molecular biology technique. However, because it is prone to amplicon and sample

contamination, planning and designing of your PCR lab space will need careful consideration.

CHAPTER 1: What you need to know about PCR

Designing your PCR lab

Ideally, a PCR lab should have two rooms with two areas, each designed for specific tasks.

The first room should be exclusively used for pre-PCR activities and divided into a master mix

preparation area and a sample preparation area. Air pressure should be slightly positive to

prevent aerosols from flowing in.

The second room should have a dedicated area for nucleic acid amplification, and another one

for product analysis. Air pressure should be slightly negative to ensure that amplicon aerosols

don't leave the room.

If you're lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible.

Having pre-PCR activities spatially separated from the amplification and analysis area – either

in different rooms or in separate benches – is very important, because you usually have a

low amount of the nucleic acid sample during preparation and a very high concentration after

CHAPTER 1: What you need to know about PCR 21

amplification. This means that if you analyze your PCR in the same space as you prepare your

master mix and samples, you may get false-positive results due to amplicon contamination.

You should also ensure that your lab set-up follows a unidirectional workflow. No materials or

reagents used in the amplification and analysis areas should ever be taken into the pre-PCR

space without a thorough decontamination. This means that you'll need dedicated equipment

for each area, e.g., two different sets of pipettes. This unidirectional workflow should also apply

to lab staff. If you've been working in the amplification and analysis areas, and you need to go

back to the pre-PCR area, change your personal protective equipment, as it may have been

contaminated by amplicon aerosols.

Another precautionary measure to take into account when setting up your PCR lab, in addition

to the spatial separation, is temporal separation. You could, for example, consider setting up

your PCR reactions in the morning, and perform the amplification and analysis steps in the

afternoon. This may limit your flexibility, but will prevent contamination issues and having to

repeat your experiment.

PCR equipment tips

PCR labs typically require a variety of equipment, such as centrifuges, vortex mixers, pipettes,

fridges and freezers, thermal cyclers and analysis instruments (e.g., electrophoresis systems).

Depending on the size of your lab and your applications, the amount of equipment you’ll need

may vary. Instead of providing you a 'shopping list', we will outline what you should look for

when purchasing equipment and consumables in order to keep contamination of your PCR

reactions to a minimum.

22 CHAPTER 1: What you need to know about PCR

Laminar flow or biosafety cabinet

Since you can never be 100 % certain that there are no amplicon aerosols in your pre-PCR

space, you should set up your PCR reactions in a laminar flow hood or biosafety cabinet,

decontaminated with a bleach solution prior to starting and after you finish your work.

Pipette tips and other consumables

Despite being more expensive than normal pipette tips, using filter tips for your PCR set-up will

avoid aerosols entering and contaminating your pipette, and avoid aerosols that might already

be present in your pipette contaminating your master mix or samples. To minimize your filter tip

consumption, first fill all your tubes with the master mix using only one tip or set of tips – if you're

using multichannel pipettes – and follow with your samples, using one tip per sample. Adding

the sample last is also recommended because it's easier to dispense it into a liquid than into an

empty tube, and because it reduces the risk of aerosolizing your sample as you pipette.

For consumables, you should make sure that you have enough small vials available in your lab

when your PCR reagents arrive. Aliquoting them into smaller containers will increase their shelf

life and prevent them from going through too many freeze/thaw rounds. If your reagents get

contaminated, it will also save you from throwing away your entire supply, as you’ll have clean

aliquots available for a second PCR.

Finally, you’ll need to make sure that all consumables and equipment are free of DNase, RNase

and PCR inhibitors. Always choose sterile products from manufacturers that can certify that

their tips and consumables are free of any of these potential contaminants.

CHAPTER 1: What you need to know about PCR 23

Cleaning and contamination control

You won’t need to worry about cleaning or contamination control when setting up your lab, but

you will when your lab is up and running. We will briefly address this topic below.

Whether you decide to set up your PCR reactions in a laminar flow hood, a biosafety cabinet

or an open bench, you will need to decontaminate your work space before and after set-up by

wiping it with a freshly made bleach solution and distilled water. The same process should be

performed in the amplification and analysis areas. You should also make sure you clean your

pipettes, equipment, doorknobs, and the handles of your fridges and freezers regularly.

Because PCR assays are so sensitive, all the preventative measures described here may still

not guarantee that your experiments will never get contaminated. It is therefore necessary

to include the appropriate controls to detect contamination early. Always include negative

and positive controls, as this will help identifying master mix contaminations, and confirm the

performance of the extraction protocol, reagents and amplification steps. Additionally, you

should monitor the positivity rate in your lab, and ensure that unexpected increases in detection

have identifiable causes, e.g., a seasonal outbreak.

Conclusion

In this article, we covered how to set up your PCR lab to ensure spatial and temporal

separation, and prevent contamination. We also outlined the key factors to consider when

purchasing equipment and consumables for your lab, to maintain safety and reduce wastage.

Lastly, we highlighted the importance of regular workspace cleaning and the use of appropriate

controls to detect any contamination early on. We hope that you are still just as excited about

setting up your PCR lab, and that this article has made the task less daunting for you.

24 CHAPTER 1: What you need to know about PCR

1.4 qPCR: How SYBR® Green and TaqMan®

real-time PCR assays work

qPCR, or real-time PCR, is a widely used method to quantify DNA sequences in samples.

This article gives you a comprehensive introduction to the topic, explaining how dye-based

and probe-based qPCR assays (like SYBR Green and TaqMan) work, how to validate your

amplification experiments, and how to analyze your qPCR data.

qPCR vs PCR vs RT-PCR

Before explaining how qPCR works, we would like to briefly outline its difference from standard

PCR and RT-PCR.

Whereas standard PCR monitors DNA amplification upon reaction completion, qPCR uses

fluorescent signals to monitor DNA amplification as the reaction progresses. This is why qPCR

is also referred to as real-time PCR, quantitative PCR or quantitative real-time PCR.

RT-PCR, not to be confused with real-time PCR, stands for reverse transcription PCR and can

be used to amplify RNA target sequences. It involves an initial incubation of the sample RNA

with a reverse transcriptase enzyme and a DNA primer before amplification.

How qPCR works

qPCR relies on fluorescence from intercalating dyes or hydrolysis probes to measure DNA

amplification after each thermal cycle. The most common dye-based method is SYBR Green,

and the most common probe-based method is TaqMan, which is why this article will focus on

these two qPCR techniques.

CHAPTER 1: What you need to know about PCR 25

SYBR Green qPCR

Like standard PCR, the SYBR Green protocol consists of denaturation, annealing and

extension phases. The difference being that you add some double-stranded DNA binding dye,

SYBR Green I, to your master mix during qPCR setup. This fluorescent dye intercalates into

double-stranded DNA sequences during the extension phase, where it shows a strong increase

in fluorescent signal. Measuring this signal at the end of every thermal cycle will allow you to

determine the quantity of double-stranded DNA present.

The downside of the SYBR Green assay is that the dye binds to any double-stranded DNA

sequence. This means that you could also detect fluorescence emitted from non-specific

qPCR products, such as primer dimers. To eliminate this risk, check the reaction specificity by

performing a melting curve analysis, explained later in the article, or use the TaqMan method.

TaqMan qPCR

Instead of using intercalating dyes, this assay uses TaqMan probes with a 5' fluorescent

reporter dye and a 3' quencher dye. These probes are target-specific, and only bind to the

DNA sequence of interest downstream of one of the primers during the annealing step. When

the enzyme Taq-polymerase encounters the TaqMan probe during the extension phase, it

displaces and cleaves the 5' reporter dye. Once the reporter dye has been separated from

the quencher dye, its measurable fluorescent signal at the end of every qPCR cycle increases

significantly. The second DNA strand is synthesized in parallel but, as no probe is attached to it,

this process can't be monitored by fluorescence measurements.

Compared to the SYBR Green assay, the use of TaqMan probes is more expensive, but also

offers two significant advantages:

• the TaqMan assay only measures amplification progression of the target sequence, as the

probes are target specific.

• you can monitor the quantity of various qPCR products in a single reaction by adding different

primers and TaqMan probes with different reporter dyes to the master mix. This multiplex

approach allows you to detect several fluorescent signals at the end of every thermal cycle.

26 CHAPTER 1: What you need to know about PCR

Amplification plot

For both qPCR methods, data is visualized in an amplification plot, with the number of thermal

cycles on the x-axis, and the fluorescent signals detected on the y-axis:

CHAPTER 1: What you need to know about PCR 27

As can be seen, fluorescence remains at background levels during the first thermal cycles.

Eventually, the fluorescent signal reaches the fluorescence threshold, where it is detectable over

the background fluorescence. The cycle number at which this happens is called the threshold

cycle (Ct). If the Ct value for a sample is high, it means that little starting material was present,

and vice versa. Please note that you should always analyze at least three replicates of each

sample, as tiny pipetting errors during qPCR set-up can result in huge differences in Ct values.

The Ct value is sometimes also referred to as crossing point (Cp), take-off point (TOP) or

quantification cycle (Cq) value, with MIQE guidelines suggesting using Cp value to standardize

terminology.1 In this article we will continue to call it Ct, as this is the most commonly used term.

MIQE guidelines

The MIQE (Minimum Information for Publication of Quantitative Real-Time PCR Experiments)

guidelines describe the minimum information necessary for evaluating qPCR experiments.

When publishing a manuscript, the scientist needs to provide all relevant experimental

conditions and assay characteristics described by the MIQE guidelines, allowing reviewers

to assess the validity of the protocols used, and enabling other scientists to reproduce the

experiments.

Validation of qPCR assays

qPCR amplification plots can be analyzed using absolute or relative quantification. However,

before explaining qPCR data analysis, we need to quickly discuss how to determine reaction

efficiency and specificity. You don't need to perform these steps after every qPCR experiment,

but should always validate these two values when setting up a new qPCR protocol or changing

your current workflows.

Reaction efficiency

A perfect qPCR assay would have a reaction efficiency of 100 %, which means that the number

of template DNA copies would double at every thermal cycle. As this is almost impossible to

achieve in practice, reaction efficiencies between 90 and 110 % are considered to be ideal.

To calculate the reaction efficiency of your assay, you need to set up a 10-fold serial dilution

of an undiluted sample with a known amount of template DNA. After running a qPCR, create a

standard curve with the log of the starting quantity on the x-axis and the Ct values on the y-axis.

28

Using the equation for the linear regression line (y = mx + b), you can now determine the

reaction efficiency as follows2:

Efficiency = (10(-1/m)-1) x 100

In our example, m would be -3.5826, resulting in a reaction efficiency of 90.1634 %.

Reaction specificity

Reaction specificity can be determined using a melting curve analysis, allowing you to identify

non-specific qPCR products and primer-dimers. To perform a melting curve analysis, run a

qPCR assay with a fluorescent intercalating dye like SYBR Green I. After amplification, the

thermal cycler increases the temperature step by step while monitoring fluorescence. As

the temperature increases, the dsDNA qPCR products present will denature, resulting in a

decreasing fluorescent signal:

CHAPTER 1: What you need to know about PCR

CHAPTER 1: What you need to know about PCR 29

Then, plot the change in slope of this curve as a function of temperature to obtain a melting

curve:

If you're observing only one melting peak like the image above, your qPCR assay is specific.

If there are several melting peaks, primer-dimers and/or non-specific products were amplified

during qPCR, and you should redesign your experiment to increase its specificity.

30 CHAPTER 1: What you need to know about PCR

Analysis of qPCR data

qPCR data can be analyzed by absolute or relative quantification, and the method suitable

for your experiment depends on your goal. Absolute quantification allows you to determine

the quantity of starting material that was present in a given sample before amplification. For

example, this method can be used to determine the viral load of a patient sample. Relative

quantification is applied to compare levels or changes in gene expression between different

samples. For example, it is helpful to investigate whether the expression of a certain gene is

higher in a tumor sample than in a healthy control sample.

Absolute quantification

After qPCR amplification, you will have produced an amplification plot, and know the Ct value

of each sample. To find the quantity of starting material present in your samples, you need to

compare these values to a standard curve. As seen above in the section on reaction efficiency,

a standard curve is obtained by amplifying a serial dilution of a sample with a known amount of

template DNA, then plotting the Ct values against the log of the starting quantities.

The equation for the linear regression line of the standard curve (y = mx + b) will then allow you

to calculate the quantity of starting material for each sample. As y corresponds to the Ct value,

and x to the log quantity, the equation for the linear regression line is equivalent to:

Ct = m(log quantity) + b

Solving this equation for the quantity will give you the formula:

Quantity = 10((Ct-b)/m)

This will allow you to quickly determine the quantity of starting material in each sample.

Y = mx + b → Ct = m(log quantity) + b → Quantity = 10((Ct-b)/m)

Relative quantification

To compare levels or changes in target gene expression between different samples and a

control sample, you first need to define a reference gene whose expression is unregulated.

Then, run a qPCR to obtain the Ct values for the reference gene, target gene in your samples,

and the control sample.

If the reaction or primer efficiencies for the reference and target genes are near 100 %, and

within 5 % of each other, you can then use the ΔΔCt method – also called the Livak method – to

determine the expression rate of the target gene in your samples. However, if the efficiencies

are further apart, you should use the Pfaffl method. To learn how to calculate reaction

efficiencies, please refer to the 'Reaction efficiency' section earlier in the article.

CHAPTER 1: What you need to know about PCR 31

The calculations for the two methods are as follows:

ΔΔCt method

Normalize the Ct of the target gene to the Ct of the reference gene for each sample and the

control sample:

ΔCt(sample) = Ct(target gene) – Ct(reference gene)

ΔCt(control) = Ct(target gene) – Ct(reference gene)

Normalize the ΔCt of each sample to the ΔCt of the control sample:

ΔΔCt(sample) = ΔCt(sample) – ΔCt(control)

Since the calculations are in logarithm base 2, you must use the following equation to get the

normalized expression ratio for each sample:

Normalized expression ratio = 2-ΔΔCt(sample)

Pfaffl method

Calculate the ΔCt of the target gene for each sample:

ΔCt(target gene) = Ct(target gene in control) – Ct(target gene in sample)

Calculate the ΔCt of the reference gene for each sample:

ΔCt(reference gene) = Ct(reference gene in control) – Ct(reference gene in sample)

Calculate the normalized expression ratio for each sample:

Normalized expression ratio = ((Etarget gene)ΔCt(target gene)) / ((Ereference gene)ΔCt(reference gene))

Etarget gene: Reaction efficiency of the target gene

Ereference gene: Reaction efficiency of the reference gene

The normalized expression ratio obtained using the ΔΔCt or the Pfaffl method is the fold

change of the target gene in your sample relative to the control. A normalized expression ratio

of 1.2 would mean that you have a gene expression of 120 % relative to the control.

Conclusion

We hope that this article answered all your questions regarding qPCR methods, assay

validation and data analysis.

32

1.5 How to design primers for PCR

PCR is one of the most widespread molecular biology applications, yet it is anything but simple

to perform. Common issues – such as a low product yield or non-specific amplification – are

often caused by poorly designed PCR primers. We have therefore summarized the most

important information on designing PCR primers to help you overcome these challenges.

What is a PCR primer?

Primers – also called oligonucleotides or oligos – are short, single-stranded nucleic acids used

in the initiation of DNA synthesis. During PCR reactions, they anneal to the plus and minus

strands of the template DNA, flanking the sequence that needs to be amplified.

How to design PCR primers?

PCR primers have to be tailored to both the region of interest of your template DNA and your

reaction conditions. This means that, unlike the other components of the PCR master mix, you

can't just buy them, but need to design them yourself using a primer design tool. These tools

allow you to set parameters such as primer length, melting temperature, GC content and more.

Read on to learn what the optimal values for each of these parameters are, and how they affect

your PCR assay.

CHAPTER 1: What you need to know about PCR

CHAPTER 1: What you need to know about PCR 33

Primer length

The optimal length of a PCR primer lies between 18 and 24 bp. Longer primers are less efficient

during the annealing step, resulting in a lower amount of PCR product. Conversely, shorter

primers are less specific during the annealing phase, leading to more non-specific binding and

amplification. However, there are exceptions to this rule. For example, some scientists have

successfully used miniprimers that are 10 bp long to expand the scope of detectable sequences

in microbial ecology assays.

Target sequence length

The target sequence to be amplified should ideally be between 100 and 3000 bp for standard

PCR assays, and 75 and 150 bp for qPCR assays. Longer sequences usually need special

enzymes and reaction conditions to ensure that they are completely and specifically amplified.

Primer melting temperature

The primer melting temperature (Tm) can be defined as the temperature at which half of the

primers dissociate from the template DNA. It is usually between 50 and 60 °C, and the melting

temperatures of the forward and reverse primers should be within 5 °C of each other. If the two

melting temperatures are further apart, it won't be possible to find an annealing temperature

that allows both primers to bind to the template DNA.

Most primer design tools use the nearest neighbor method to calculate primer melting

temperatures, as it's the most accurate. However, if you want to make an approximate

calculation yourself, you can use this formula:

Tm = 4 °C x (G+C) + 2 °C x (A+T)

Tm: melting temperature

G, C, A, T: number of nucleobases (guanine, cytosine, adenine, thymine) in the primer

As indicated in the formula above, G-C bonds are harder to break than A-T bonds – because

G-C base pairs are linked by three hydrogen bonds, and A-T base pairs by two – and the length

of the primer also impacts its melting temperature. This means that you can either increase the

GC content of a primer (provided the template allows for this), or slightly extend its length if its

melting temperature is too low.

34

Primer annealing temperature

The primer annealing temperature (Ta) is the temperature needed for the annealing step of

the PCR reaction to allow the primers to bind to the template DNA. The theoretical annealing

temperature can be calculated as follows:

Ta = 0.3 x Tm(primer) + 0.7 x Tm(product) – 14.9

Ta: primer annealing temperature

Tm(primer): lower melting temperature of the primer pair

Tm(product): melting temperature of the PCR product

Once you've calculated the theoretical annealing temperature, the optimal annealing

temperature needs to be determined empirically. To achieve this, perform a gradient PCR,

starting a few degrees below the calculated annealing temperature, and ending a few degrees

above. After amplification, run a gel, and the sample producing the clearest band contains the

largest quantity of PCR product, making its annealing temperature the optimal one for your

primers. Usually, you'll get a value that is 5 to 10 °C lower than the primer melting temperature.

CHAPTER 1: What you need to know about PCR

35

It's important to determine the optimal annealing temperature, as primers could form hairpins or

bind to regions outside the DNA sequence of interest if it's too low, producing non-specific and

inaccurate PCR products. If the annealing temperature is too high, the primers won't sufficiently

bind to the template DNA, and you'll obtain little to zero amplicons.

CHAPTER 1: What you need to know about PCR

GC content

As seen before, G-C base pairs are stronger than A-T base pairs, which means that a higher

GC content ensures a more stable binding between the primers and the template DNA. The

optimal GC content of a primer lies between 40 and 60 %, and primers should have two to three

Gs and Cs at the 3' end to bind more specifically to the template DNA.

Runs and repeats

Avoid runs of four or more single bases – such as ACCCCC – or dinucleotide repeats (for

example, ATATATATAT), as they can cause mispriming.

Cross homology

If a primer is homologous to a template DNA sequence outside the region of interest, these

sequences will be amplified too. Therefore, you should always test the specificity of your

primer design against genetic databases; for example, by ‘blasting’ them through NCBI BLAST

software.

36

Your PCR product yield will be less if secondary structures form and remain stable above the

annealing temperature of your reaction, as the primers bind to themselves or another primer

instead of the template DNA. This is why your primer design tool should be able to check for,

and warn you of stable secondary structures.

Mismatches and degenerated positions

Mismatches are primer bases that aren't complementary to the target sequence. They can be

tolerated to a certain extent, and are sometimes even necessary; for example, when performing

a multi-template PCR to amplify a set of similar target sequences from different bacteria with

a single set of primers. Degenerate primers could help if mismatches negatively impact the

performance of your PCR.

Degenerate primers have several different nucleotides in some of their positions. For example,

instead of A you could have an equal concentration of A and T in a certain position. The codes

for the different nucleotide combinations available for degenerate primers are as follows:

CHAPTER 1: What you need to know about PCR

Secondary structures

There are three different types of secondary structures – also called primer dimers – that can

form during a PCR assay:

• Hairpins: caused by intra-primer homology – when a region of three or more bases is

complementary to another region within the same primer – or when a primer melting

temperature is lower than the annealing temperature of the reaction.

• Self-dimers: formed when two same sense primers have complementary sequences – interprimer

homology – and anneal to each other.

• Cross-dimers: formed when forward and reverse primers anneal to each other when there is

inter-primer homology.

37

IUPAC NUCLEOTIDE CODE BASE

R A or G

Y C or T

S G or C

W A or T

K G or T

M A or C

B C or G or T

D A or G or T

H A or C or T

V A or C or G

N Any base

Conclusion

This article summarized the key points to consider when designing PCR primers to help avoid

common issues like low product yield or non-specific amplification. We covered optimal primer

and target sequence lengths, and ideal primer melting and annealing temperatures. We also

provided helpful tips for other crucial factors such as GC content, runs and repeats, cross

homology and the danger of stable secondary structures. Lastly, the article highlighted the

value and pitfalls of mismatches and degenerated positions. That's it, after reading about all of

this, you are sure to be a 'PCR Primer Pro'!

CHAPTER 1: What you need to know about PCR

38

CHAPTER 2:

INTEGRA Biosciences’ PCR solutions

PCR is a robust method, but it’s comprised of numerous stages, involving multiple precise

pipetting steps that often prove time consuming and prone to errors. Temperature-sensitive

reagents and samples may affect accuracy, and the varying viscosities of samples, as well as

‘sticky’ DNA, can be difficult to handle. On top of this, the repetitive nature of this work can also

frequently result in user fatigue and handling mistakes.

Fortunately, the right tools can eliminate your pipetting predicaments, vastly improving the

reproducibility and productivity of your PCR workflows. Here, we will demonstrate how our

range of liquid handling solutions are perfect for PCR applications, allowing you to create a

faster and more efficient workflow with fewer errors.

Manual and electronic pipettes

A good starting point for lower throughput PCR applications – up to half a plate per day – is our

EVOLVE single or multichannel pipettes, which feature convenient volume adjustment dials to

increase the accuracy and speed of manual handling. Our range of VIAFLO electronic pipettes

is also suitable for low throughput PCR set-up, and can easily handle up to eight plates per day.

CHAPTER 2: INTEGRA Biosciences’ PCR solutions

Learn more

about

EVOLVE

Learn more

about

VIAFLO

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 39

Learn more

about

VOYAGER

Adjustable tip spacing pipettes

PCR set-up usually requires transferring liquids between different labware formats which is

tedious and highly error prone. Our VOYAGER adjustable tip spacing pipettes solve these

problems, increasing speed and eliminating transfer errors, while ergonomic single-handed

operation leaves the other hand free to handle labware.

96 and 384 channel pipettes

We have a wide range of options perfect for productive high throughput PCR set-up – more

than eight plates per day – which are suitable for different lab sizes and budgets. Our

VIAFLO 96 and VIAFLO 384 channel handheld electronic pipettes, as well as the

MINI 96 channel portable electronic pipette, can reduce handling steps while

increasing productivity and reproducibility.

Learn more

about

MINI 96

Learn more

about

VIAFLO 96

and VIAFLO 384

40 CHAPTER 2: INTEGRA Biosciences’ PCR solutions

Pipetting robots

INTEGRA also offers pipetting robots for high throughput laboratories, or for labs that want to

reduce the risk of contamination due to manual processing. For example, the ASSIST PLUS

pipetting robot can automate the D-ONE single channel pipetting module for master mix

preparation, and VIAFLO and VOYAGER multichannel pipettes to take care of the multiple

pipetting steps in PCR workflows.

Learn more

about

D-ONE

Learn more

about

GRIPTIPS

Learn more

about

ASSIST PLUS

Pipette tips

INTEGRA has developed GRIPTIPS pipette tips to complement its range of pipetting solutions.

GRIPTIPS are free from RNase, DNase and PCR inhibitors, and perfectly fit all INTEGRA

pipetting solutions, reducing the risk of contamination from tips that leak or fall off.

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 41

Learn more about

sample transfers

from plate to plate

Learn more about

sample transfers

from tubes to plates

Sample reformatting

The transfer of samples between different labware formats is a slow, tedious and highly

error-prone procedure when performed manually with a single channel pipette. The

combination of the ASSIST PLUS pipetting robot and VOYAGER adjustable tip spacing

pipette provide a novel solution for automated, accurate and efficient liquid transfer of multiple

samples in parallel. For even higher throughput applications, the VIAFLO 96, VIAFLO 384

and MINI 96 offer a fast solution for whole plate transfers.

42 CHAPTER 3: Application Notes

CHAPTER 3:

Application notes

Our pipetting instruments are used across a broad spectrum of life sciences applications. To

help share this knowledge and experience of using INTEGRA products with the wider scientific

community, we have developed an application database which contains a wide range of useful

application notes. Here are some of the most relevant app notes related to PCR protocols and

workflows.

3.1 Efficient and automated 384 well qPCR set-up

with the ASSIST PLUS pipetting robot

Using the ASSIST PLUS pipetting robot to automate set-up for a 384 well

plate qPCR

Setting up a qPCR is a tedious process consisting of multiple pipetting steps. One particularly

challenging task is reformatting from microcentrifuge tubes into a 384 well plate, which is time

consuming and requires a lot of concentration. Another common problem is the loss of valuable

and expensive substances, such as master mix and

precious samples, due to the reservoir dead volume.

The ASSIST PLUS pipetting robot, in combination with

the VIAFLO and VOYAGER electronic pipettes,

streamlines the workflow and increases the throughput

and the reproducibility of qPCR set-ups, with minimal

manual input. The loss of expensive substances or

valuable samples due to reformatting errors is

eliminated. The unique design of the ASSIST PLUS

pipetting robot, together with the intuitive

VIALAB software, offers

exceptional flexibility and

straightforward implementation.

CHAPTER 3: Application Notes 43

Key benefits

• Automating the qPCR set-up with the

VIAFLO 16 channel electronic pipette and

the ASSIST PLUS pipetting robot allows

considerably faster sample preparation,

freeing up time for scientists to focus on

other experiments.

• Automation of VOYAGER adjustable tip

spacing pipettes with the ASSIST PLUS

offers a reliable pipetting method that

requires minimal manual intervention and

eliminates the risk of reformatting errors.

• The use of low retention GRIPTIPS with

heightened hydrophobic properties and

SureFlo™ low dead volume reservoirs with

an anti-sealing array helps to save precious

samples and master mix. Combined with

the high pipetting accuracy and precision

of the ASSIST PLUS pipetting robot, this

enables exceptionally low dead volumes to

be achieved.

• The ASSIST PLUS pipetting robot, in

combination with the intuitive VIALAB

software, is quick to set up and easy to use.

Overview: qPCR set-up

The ASSIST PLUS pipetting robot is used to set up a 384 well format qPCR by pipetting 64

samples in triplicate with two different master mixes for the detection of two genes of interest

(GOI 1 and GOI 2).

The protocol is divided into two programs that guide the user through all the steps of the qPCR

set-up:

• Program 1: Mastermix_qPCR

• Program 2: Samples_qPCR

The ASSIST PLUS pipetting robot operates a VIAFLO 16 channel 125 μl electronic pipette with

125 μl sterile, filter, low retention GRIPTIPS for program 1 and a VOYAGER 8 channel 12.5 μl

electronic pipette with 12.5 μl sterile, filter, low retention GRIPTIPS for program 2.

44

Experimental set-up: Program 1 - master mix

transfer (Mastermix_qPCR)

Prepare the pipetting robot deck as follows (Figure 1):

Deck position A: Dual reservoir adapter – 2 x 10 ml reagent

reservoir with SureFlo anti-sealing array

(Figure 2) containing master mix 1 and 2.

Deck position B: 384 well PCR plate, placed on an INTEGRA

cooling block in the landscape position.

CHAPTER 3: Application Notes

Figure 1: Set-up for the master mix transfer. Position A: dual reservoir adapter with 2 x 10 ml reagent

reservoirs with SureFlo anti-sealing array. Position B: 384 well PCR plate, placed on an INTEGRA

cooling block.

Figure 2: The INTEGRA dual reservoir adapter accommodates both 10 ml reagent reservoirs on one

deck position.

CHAPTER 3: Application Notes 45

Step-by-step procedure

1. Transfer master mixes into the 384 well plate

Add master mixes 1 and 2 into the left and right sides of the 384 well PCR

plate, respectively.

Use an EVOLVE 5000 μl manual pipette with

5000 μl sterile, filter, low retention GRIPTIPS to

fill the left 10 ml reagent reservoir with SureFlo

anti-sealing array with 1.6 ml of master mix 1 and

the right reservoir with 1.6 ml of master mix 2

(position A). Select and run the VIALAB program

‘Mastermix_qPCR’ on the VIAFLO 16 channel

125 μl electronic pipette with 125 μl sterile, filter,

low retention GRIPTIPS. The ASSIST PLUS

pipetting robot automatically transfers 7.5 μl of

master mix 1 (pink) into the left half of the 384 well

PCR plate and 7.5 μl of master mix 2 (blue) into the

right half (Figure 3) using the Repeat Dispense

mode with a tip touch on the surface of the liquid to

increase pipetting precision. Figure 4 shows the

pipetting robot transferring the master mix into a

384 well plate.

Tips:

• Pre- and post-dispense steps are recommended

to increase the accuracy and precision of

pipetting. The pre- and post-dispense volumes

should be between 3 and 5 % of the nominal

volume of the pipette.

• The low retention GRIPTIPS are made from a

unique polypropylene blend with heightened

hydrophobic properties for superior accuracy

and precision while pipetting viscous and low

surface tension liquids.

• The reservoirs’ SureFlo anti-sealing array and

a unique surface treatment that spreads liquid

evenly enable the pipette tips to sit on the bottom

and still aspirate liquids accurately, reducing

dead volumes.

Figure 3: Pipetting scheme for master mixes 1 (pink) and 2 (blue).

Figure 4: Example of the ASSIST PLUS pipetting robot

transferring a master mix into a 384 well PCR plate.

46 CHAPTER 3: Application Notes

Experimental set-up: Program 2 - sample transfer

(Samples_qPCR)

Prepare the pipetting robot deck as follows (Figure 5):

Deck position B: 384 well PCR plate, placed on an INTEGRA

cooling block.

Deck position C: INTEGRA 1.5 ml microcentrifuge tube rack,

with tubes containing samples 1-32.

Figure 5: Set-up for the sample transfer protocol. Position B: 384 well PCR plate, placed on an INTEGRA

cooling block. Position C: INTEGRA 1.5 ml microcentrifuge tube rack, with tubes containing samples 1-32

(Figure 6).

Figure 6: Example of the ASSIST PLUS pipetting samples from the INTEGRA microcentrifuge tube rack

into a 96 well plate.

VOYAGER - 12.5 μl – 8CH

12.5 μl GRIPTIP,

sterile, filter

B 384 well PCR Sapphire on 384 well cooling block – 45 μl C Rack for 1.5 ml microcentrifuge tubes – 1500 μl

CHAPTER 3: Application Notes 47

Step-by-step procedure

1. Sample transfer into the 384 well plate

Add the 64 samples in triplicate to the master mixes.

Place samples 1-32 in an INTEGRA 1.5 ml microcentrifuge tube rack on position C. Run the

VIALAB program ‘Samples_qPCR’ on a VOYAGER 8 channel 12.5 μl electronic pipette to start

the sample transfer. The ASSIST PLUS transfers 2.5 μl of the first 32 samples in triplicate into

master mixes 1 and 2 (Figure 7, yellow/brown), using the Repeat Dispense mode with a tip

touch on the side of the well to make sure that no droplets adhere to the GRIPTIPS. After this

step, a prompt informs the user to place the second series of samples (33-64) on position C.

The ASSIST PLUS pipetting robot continues by transferring 2.5 μl of the samples in triplicate

into the other half of master mixes 1 and 2 (Figure 7, green).

Tip: Use sterile, filter, low retention GRIPTIPS for optimal liquid recovery of precious solutions,

such as the master mix and samples.

Figure 7: Pipetting scheme of the qPCR assay.

master mix 1 master mix 2

Sample

33 - 64

Sample

1 - 32

48 CHAPTER 3: Application Notes

Remarks

VIALAB software:

The VIALAB program can easily be adapted to fit the user’s demands, especially if specific

labware, incubation times or protocols are needed.

Partial plates:

The programs can be adapted at any time to a different number of samples, giving

laboratories total flexibility to meet current and future demands.

Conclusion

• The time required for a 384 well qPCR set-up can be reduced from 1.5 hours using

a single channel pipette to 12 minutes using the ASSIST PLUS pipetting robot in

combination with VIAFLO 16 channel and VOYAGER 8 channel pipettes.

• The ASSIST PLUS, together with the VOYAGER adjustable tip spacing pipette,

guarantees perfectly reproducible test results and eliminates all risks of reformatting

errors when transferring samples from microcentrifuge tubes into a 384 well plate.

• INTEGRA’s low retention GRIPTIPS increase pipetting precision for viscous or low

surface tension liquids. The reagent reservoirs with SureFlo anti-sealing array reduce the

dead volume of costly reagents and precious samples.

• The intuitive VIALAB qPCR program is quick to set up and easy to use or adapt to other

pipetting protocols.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 3: Application Notes 49

3.2 Automated RT-PCR set-up for

COVID-19 testing

How to prepare RT-PCR plates for SARS-CoV-2 detection

with the ASSIST PLUS

The emergence and outbreak of the novel coronavirus

SARS-CoV-2 (COVID-19) has placed unprecedented

demands on laboratories testing for COVID-19, leaving

scientific staff to contend with a spiraling influx of patient

samples and a rapid, continuous growth in workload.

Laboratories need additional automated liquid handling

instruments for viral nucleic acid extraction

and RT-PCR set-up – which are the

most labor-intensive processes in

the diagnostic testing workflow – to

increase the sample throughput

capacity, reduce manual

intervention by laboratory

analysts and fast track the

development of COVID-19 assays.

The ASSIST PLUS pipetting robot together with a VOYAGER

adjustable tip spacing pipette, low retention GRIPTIPS and SureFlo

10 ml reagent reservoirs were successfully used for RT-PCR set-up in

COVID-19 testing laboratories.

50 CHAPTER 3: Application Notes

Key benefits

• The full automation capability of the

ASSIST PLUS reduces manual intervention

and frees highly valuable time for laboratory

personnel in this critical COVID-19

pandemic.

• The compact and easy-to-use

ASSIST PLUS pipetting robot allows

fast set-up regarding installation and

programming, allowing labs to immediately

increase their sample processing capacity

and fast track assay development for

COVID-19 sample testing.

• VOYAGER adjustable tip spacing pipettes

in combination with the ASSIST PLUS

provide unmatched pipetting ergonomics by

automatically reformatting patient samples

from tube racks into 384 well plates.

• Optimal pipette settings, including tip

immersion depth, pipetting speeds and

angles, deliver reproducible, precise and

accurate results, with no contamination

observed in controls or patient samples.

• The use of INTEGRA’s low dead volume,

SureFlo 10 ml reagent reservoirs, together

with low retention GRIPTIPS, demonstrated

excellent results, enabling efficient handling

of the precious and expensive one-step RTPCR

master mix used for patient testing.

Overview: Automated RT-PCR set-up

The ASSIST PLUS pipetting robot is used to automate testing of suspected COVID-19

positive cases in a 384 well plate. The pipetting robot operates a VOYAGER 12 channel 50 μl

electronic pipette with 125 μl sterile, filter, low retention GRIPTIPS. To double the available

testing capacity and, concurrently, decrease the cost per test of expensive one-step RT-PCR

reagents of dwindling availability, the total PCR reaction volume was miniaturized, reducing it

to 10 μl – inclusive of 7.5 μl one-step RT-PCR master mix and 2.5 μl of nucleic acid template.

The templates were extracted from combined nasopharyngeal/oropharyngeal flocked swabs

or sputum samples. The following procedure is based on the protocol used by the Microbiology

and Molecular Pathology Department at Sullivan Nicolaides Pathology (SNP) – part of the

Sonic Healthcare Group – in Brisbane, Australia.

The protocol is divided into two parts:

• Program 1: Add the master mix (1-COVID-19)

• Program 2: Add the nucleic acid template (2-COVID-19)

CHAPTER 3: Application Notes 51

Experimental set-up: Program 1

Deck position A: 10 ml reagent reservoir with

SureFlo anti-sealing array containing

3 ml of one-step RT-PCR master mix.

Deck position C: 384 well plate placed on a PCR 384 well

cooling block, allowing the master mix and

samples to be kept cold, and enabling exact

positioning of the PCR plate on the deck.

Figure 1: The set-up for program 1-COVID-19.

VOYAGER - 50 μl – 12CH

50/125 μl GRIPTIP, sterile, filter,

low retention

A Multichannel reservoir – 10ml C PCR cooling block 384_system

52 CHAPTER 3: Application Notes

Step-by-step procedure

1. Add the master mix

Fill the 384 well plate with the one-step RT-PCR master mix.

Place the one-step RT-PCR master mix in a 10 ml sterile, polystyrene reagent reservoir with

INTEGRA’s SureFlo anti-sealing array. Set up the deck with the required labware, as indicated

in Figure 1. Select the VIALAB program 1-COVID-19. The VOYAGER pipette automatically

transfers the master mix from the reservoir into the 384 well plate (LightCycler® 480 Multiwell

Plate, Roche) using the Repeat Dispense mode with tip touch. Each well of the plate is filled

with 7.5 μl of master mix.

Tips:

• Using a 10 ml reagent reservoir with SureFlo anti-sealing array allows a very low dead

volume (<20 μl) to minimize the loss of expensive reagent of dwindling availability

(see Figure 2).

• The combination of a low pipetting speed – set at 2 – and low retention GRIPTIPS shows

excellent results when pipetting the viscous and foamy master mix.

• Pre- and post-dispense settings, together with the tip touch option, guarantee reproducible,

precise and accurate pipetting results (see Figure 2).

• The PCR cooling block is used as a support to fix the position of the 384 well plate on the

deck, ensuring exact tip positioning when pipetting. The cooling block also helps to keep

samples and reagents cool if required by the protocol.

Figure 2: Precise and accurate dispensing of one-step RT-PCR master mix from the low dead

volume reagent reservoir to the 384 well plate.

CHAPTER 3: Application Notes 53

Experimental set-up: Program 2

Deck position A and B: FluidX Cluster 0.7 ml tubes containing the

nucleic acid templates. The tubes are stored

in a 96-format rack. A total of four sample

racks are used for the protocol (two on

position A and two on position B).

Deck position C: 384 well plate placed on a PCR 384 well

cooling block.

Figure 3: The set-up for program 2-COVID-19.

2. Add the nucleic acid templates

Transfer the samples from four 96-format tube racks to the 384 well plate.

Nucleic acid templates extracted from combined nasopharyngeal/oropharyngeal flocked

swabs or sputum samples are stored in FluidX Cluster 0.7 ml tubes placed in a 96-format

rack. The VOYAGER pipette transfers 2.5 μl of template from the tubes to the 384 well plate,

automatically changing the GRIPTIP pipette tips after each dispense. Both position A and B

are used to house the samples on the deck (see Figure 3). The pipette prompts the user when

it is time to replace the tube racks on the deck. After user confirmation, the VOYAGER pipette

continues reformatting the samples from tubes to the plate.

50/125 μl GRIPTIP, sterile, filter,

low retention

VOYAGER - 50 μl – 12CH

A FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl

B FluidX 96-formal, 0.7 ml Internal Thread Tube, V-Bottom

– 700 μl C PCR cooling block 384_system

54

Tips:

• The VOYAGER pipette’s tip spacing capability combined with automatic Tip Change ensures

easy and rapid sample transfer without risk of contamination or reformatting errors.

• Using an air gap of 1.5 μl when aspirating the viral nucleic acid template eliminates the risk of

contamination risk during pipette tip travel.

Note: Automated RT-PCR testing for COVID-19 with the ASSIST PLUS can also be

performed using a VOYAGER 8 channel 50 μl electronic pipette (see Figure 5).

Figure 4: Easy and rapid transfer of patient nucleic acid templates from the tube rack to the 384 well

plate using the VOYAGER adjustable tip spacing pipette together with the ASSIST PLUS pipetting robot.

Figure 5: Automated RT-PCR testing for COVID-19 using the ASSIST PLUS pipetting robot together with

a VOYAGER 8 channel adjustable tip spacing pipette, as performed in the Microbiology and Molecular

Pathology Department at SNP.

CHAPTER 3: Application Notes

55

Remarks

4 Position Portrait Deck:

If your process allows, the protocol can be compiled into one simple program using the

4 Position Portrait Deck option on the ASSIST PLUS (see Figure 6).

96 well plates:

The protocol can be readily adapted to 96 well format.

VIALAB software:

The VIALAB programs can be easily adapted to your specific labware and protocols.

CHAPTER 3: Application Notes

Figure 6: Example set-up of the 4 Position Portrait Deck when combining programs 1-COVID-19 and

2-COVID-19 in one program.

50/125 μl GRIPTIP, sterile, filter,

low retention

VOYAGER - 50 μl – 12CH

A Multichannel reservoir – 10ml B FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl

C FluidX 96-formal, 0.7 ml Internal

Thread Tube, V-Bottom – 700 μl D PCR cooling block 384_system

56

Conclusion

• In the context of a global pandemic where laboratories are under increasing pressure to

analyze more and more patient specimens to confirm COVID-19 cases, testing labs can

rapidly benefit from the advantages of the ASSIST PLUS pipetting robot, allowing them to

increase their sample processing capacity.

• Pipetting results were reproducible, precise and accurate, with no contamination

observed in controls or patient samples.

• The ASSIST PLUS pipetting robot, together with the VOYAGER adjustable tip spacing

pipette, increases sample processing capacity, reduces the need for manual intervention

by laboratory personnel and fast tracks assay development for COVID-19 testing.

• Low retention GRIPTIPS and a low dead volume SureFlo reagent reservoir allow the loss

of costly reagents, such as one-step RT-PCR master mix, to be reduced.

• The simple and fast ASSIST PLUS pipetting robot combined with the easy-to-use

VIALAB software, offers immediate help for testing labs.

• While the current protocol uses 384 well plates, it can be readily adapted to 96 well format

to meet future needs.

• Thanks to the VIALAB software, the pipetting programs can be easily adapted to any

specific protocols and labware.

CHAPTER 3: Application Notes

For more information

and a list of materials

used, please refer to

our website.

57

3.3 Increase your sample screening and genotyping

assay throughput with the VOYAGER adjustable

tip spacing pipette

Discover the advantages of setting up a genotyping assay

or sample screening with the VOYAGER adjustable

tip spacing pipette

Laboratories are continually facing the challenge of

increasing throughput in the most efficient and economical

way, to meet the need to process more and more samples

per day. Traditionally, handling and manipulating samples

between different labware formats involves the use of single

channel pipettes, especially in screening applications and

genotyping assays, which is slow, inefficient and error prone.

INTEGRA’s VOYAGER adjustable tip spacing pipette has enabled

scientists from the Technical University of Munich (TUM) to benefit

from the enhanced productivity of a multichannel pipette, reducing

tedious liquid handling tasks.

Compared to fully automated solutions, it provides seamless liquid

transfers between different standardized and non-standardized microplates,

tube and gel chamber formats, and can be used without any special training.

Tip spacing can be simply changed one-handedly with the push of a button,

eliminating the need for any manual adjustments.

The various operating modes of the VOYAGER adjustable tip spacing pipette help to speed

up monotonous pipetting steps, eliminate sample transfer errors between different labware

formats, and reduce the risk of developing repetitive strain injuries.

CHAPTER 3: Application Notes

58 CHAPTER 3: Application Notes

Key benefits

• The VOYAGER’s motorized adjustable tip

spacing enables the user to benefit from

the enhanced productivity of an electronic

multichannel pipette throughout the entire

genotyping assay, processing samples

faster than with traditional single channel

pipettes and helping to eliminate sample

transfer errors between different labware

formats.

• Tip spacing can be adjusted on the fly with

the push of a button to match different

types of labware, allowing the easy transfer

of multiple reaction mix samples from

microcentrifuge tubes directly to 96 or

384 well plates, and gel pockets.

• The availability of a range of pipetting

modes makes the VOYAGER a very

versatile and affordable tool to speed up

and standardize pipetting protocols.

• New users quickly get accustomed to the

electronic pipette thanks to its intuitive

design and easy-to-use pipetting modes.

Experimental set-up

In this protocol, two VOYAGER 8 channel adjustable tip spacing pipettes are used for a

genotyping set-up. The genotyping assay is based on a PCR method with a subsequent gel

electrophoresis.

The following protocol consists of sample transfers from 1.5 ml microcentrifuge tubes into a

96 well plate, and from a 96 well PCR plate into an agarose gel for electrophoresis.

Overview of the steps:

1. Template transfer

2. Sample transfer into the agarose gel

CHAPTER 3: Application Notes 59

Figure 1: Adjust the tip spacing by aligning it against the empty 96 well plate and tube rack.

Step-by-step procedure

1. Template transfer

Transfer the templates into a 96 well plate.

Use a VOYAGER 8 channel 300 μl electronic pipette

with 300 μl sterile, filter GRIPTIPS. Select ‘Tip spacing’

in the main menu of the pipette to set the required

spacing. Choose ‘Positions: 2’ in the tip spacing menu

and set the tip spacing according to the 96 well plate

and the microcentrifuge tubes in the rack (Figure 1).

Once saved, the tip spacing is available at any time, for

any other pipetting modes.

After saving the tip spacing, select ‘Pipet’ mode in the

main menu. Set your required sample transfer volume

and pipette the templates from the 1.5 ml microcentrifuge tubes into the 96 well plate (Figure

2). By pressing left and right on the Touch Wheel interface, the tip spacing can be adjusted on

the fly to fit each labware format.

Tips:

• Use the Repeat Dispense mode to dispense several samples successively if duplicate or

triplicate samples are required.

• Use the Pipet/Mix mode if samples require mixing in the target wells. Settings like mixing

cycles, pipetting speeds and volumes can quickly be adjusted.

Figure 2: Sample transfer from a microcentrifuge tube rack

to a 96 well plate.

60

2. Sample transfer into the agarose gel

Transfer the PCR product into the agarose gel.

After PCR, use the VOYAGER 8 channel 125 μl

electronic pipette with 125 μl sterile, filter GRIPTIPS to

transfer the samples from the 96 well PCR plate into the

agarose gel for subsequent gel electrophoresis (Figure

3). As in step 1, choose ‘Positions: 2’ in the tip spacing

menu and set the tip spacing according to the 96 well

PCR plate and the agarose gel.

Set the required sample volume as described in step 1

and transfer the samples from the PCR plate into the

agarose gel.

Tips:

• A low dispensing speed (e.g. 4) helps uniform filling of the wells in the agarose gel.

• If you want a controlled blowin – rather than automatic – keep the run button pressed while

dispensing. Blowin will occur when the run button is released.

CHAPTER 3: Application Notes

Figure 3: PCR product transfer into the agarose gel.

Conclusion

• The VOYAGER adjustable tip spacing pipette has enabled TUM researchers using

different labware formats to benefit greatly from the enhanced productivity of a

multichannel pipette, processing assays much faster than using a single channel pipette.

The tip spacing can be changed onehandedly at the touch of a button to fit different

labware formats, such as PCR plates, tubes and gel pockets.

• Thanks to the intuitive interface, users quickly become accustomed to the electronic

pipette. The different pipetting modes make the VOYAGER adjustable tip spacing pipette

a versatile yet affordable tool for working with labware of varying sizes and formats.

• The VOYAGER adjustable tip spacing pipette increases the speed of sample testing

set-ups, and helps eliminate sample transfer errors between different labware formats

and reduce the risk of developing repetitive strain injuries.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 3: Application Notes 61

3.4 PCR product purification with QIAquick® 96

PCR Purification Kit and the VIAFLO 96

handheld electronic pipette

Semi-automated PCR product purification on the

VIAFLO 96 handheld electronic pipette

QIAquick 96 PCR Purification Kit is suitable for purifying

up to 10 μg of material for downstream applications,

such as sequencing, cloning, labeling and microarrays.

The kit facilitates the removal of impurities like primers,

unincorporated nucleotides, buffers, salts, mineral oils,

agarose and enzymes. The vacuum-driven process is

much faster than centrifugation, and gives high,

reproducible yields. It is important to avoid

cross-contamination in nucleic acid purification,

and QIAGEN's column design is optimized to limit

carryover of contaminants. Although QIAquick 96

provides a high throughput solution, the elution,

washing and binding steps are very laborious and

time consuming if performed manually. With

VIAFLO 96 handheld electronic pipette, the hands-on time

is reduced, as samples and reagents can be transferred to

all 96 wells at once. This enables rapid and efficient, high throughput PCR clean-up.

Key benefits

• VIAFLO 96 and VIAFLO 384 allow

simultaneous pipetting of up to 96 or 384

wells, respectively, maximize throughput

of PCR purification by allowing transfer

samples and reagents in a single step.

• The z-heights can be predefined, choosing

the optimal value to prevent accidental

scratching of the well membrane for more

consistent results.

• Custom programming of the PCR product

clean-up steps allows pipetting parameters,

such as aspiration or dispensing speeds, to

be predefined. Prompt messages guide the

user through the entire pipetting protocol,

which is especially useful when several

pre-wetting steps are included.

• The VIAFLO 96 or VIAFLO 384's handsfree

automatic mode ensures that the

PCR clean up protocols are performed

in the same way each time, maximizing

reproducibility.

62 CHAPTER 3: Application Notes

Overview: How to purify PCR products with

VIAFLO 96

Experimental set-up

This protocol describes how PCR products

are purified using a VIAFLO 96 handheld

electronic pipette with a two position stage and

the QIAGEN QIAquick® 96 PCR Purification

Kit. The following procedure is based on the kit

manufacturer's protocol for purification of 96

samples (up to 10 μg PCR products).

A 96 channel pipetting head (50-1250 μl) is

used together with 1250 μl short, low retention,

sterile, filter GRIPTIPS. Customized VIALINK

programs are provided to perform the binding,

washing and elution steps. Before starting,

ethanol (96-100 %) should be added to the

Buffer PE concentrate.

Overview of the purification steps:

1. Step 1: Binding

2. Step 2: Washing

3. Step 3: Elution

The initial set-up of the QIAvac 96 Vacuum Manifold consists of a waste tray on top of a QIAvac

base, followed by a QIAquick 96 well plate (pink) mounted on a QIAvac 96 top plate, as shown in

Figure 1.

The QIAvac has to be attached to a vacuum source (house vacuum or vacuum pump) that

generates negative pressure between 100 and 600 mbar.

Figure 1: Initial set-up of the vacuum manifold.

CHAPTER 3: Application Notes 63

Step-by-step procedure

1. Binding

Binding the DNA to the silica-gel membrane.

Load the 1250 μl short, low retention, sterile, filter GRIPTIPS

on the VIAFLO 96. Place a 150 ml automation friendly reagent

reservoir in position A. The QIAvac 96 Vacuum Manifold

should be placed on position B of the VIAFLO 96 in landscape

orientation. No plateholder is needed on position B where the

manifold is placed.

Important: The vacuum manifold should be aligned before

each run (Figure 2).

Begin by launching the custom VIALINK program 'Qiaquick_

purification_M'. The pipette will prompt the user to place Buffer

PM on position A, then air is aspirated. This ensures that every

single drop of the liquid can be dispensed later. The

VIAFLO 96 will then guide the user through the two pre-wetting

steps, starting with aspiration and dispensing 200 μl of Buffer

PM. After a second aspiration, the pipette will display the prompt

'Move the head out of buffer', before dispensing the final 200 μl of

Buffer PM. This is followed by a 20 second wait, giving the buffer

residues time to flow down to the tip and be dispensed.

After pre-wetting, the pipette aspirates 75 μl Buffer

PM (three times the volume of the PCR product).

The instrument then tells the user to remove the

reservoir from position A, and replace it with the

96 well plate containing the 25 μl of PCR products.

After dispensing, and four mixing steps, the

resulting mixture is transferred to the QIAquick plate

wells in two steps. It is then time to switch on the

vacuum source, as indicated by the pipette.

Tips:

• Pre-wetting the tips prior to pipetting prevents

droplets and dripping when pipetting volatile

liquids, such as isopropanol, which is one of the

constituents in Buffer PM.

• Low retention GRIPTIPS (Figure 3) are used for these pipetting steps to avoid dripping.

Figure 2: Alignment of the QIAvac 96 Vacuum

Manifold.

Figure 3: Low retention versus standard tips.

64 CHAPTER 3: Application Notes

2. Washing

Two-step purification of the PCR product.

Eject the used tips and load new 1250 μl short, low retention, sterile, filter GRIPTIPS on the

VIAFLO 96. Place a new 300 ml automation friendly reagent reservoir in position A. The

VIAFLO 96 will then prompt the user to pour Buffer PE into the reservoir, followed by a prewetting

step, which is necessary since the buffer contains ethanol. After pre-wetting, the pipette

will aspirate 900 μl of Buffer PE, and dispense it into QIAquick plate wells. The instrument will

then notify the user that is it time to turn on the vacuum pump. With the pump turned on, another

dose of the buffer is dispensed into the wells, followed by a 10 minute wait to dry the membrane

and remove all residual ethanol.

Important: The final drying step is crucial to remove residual ethanol prior to elution.

Residual ethanol in the elution buffer could inhibit downstream applications (e.g. PCR).

Tip: After this step, the manufacturer suggests tapping the plate on a stack of absorbent paper

to ensure that all residual buffer is removed.

3. Elution

Elution of DNA from the silica-gel membrane.

When prompted, start by replacing the waste tray with the

blue collection microtube rack provided, which contains

1.2 ml vessels (Figure 4a). Load new 1250 μl short, low

retention, sterile, filter GRIPTIPS, and place a new

150 ml automation friendly reagent reservoir in position A.

The instrument will then prompt the user to place Buffer

EB into the reservoir, aspirate 80 μl, and dispense it into

the QIAquick plate wells. After a 1 minute incubation, the

pipette tells the user to switch on the vacuum source for

5 minutes.

Tips:

• The purified PCR product could also be eluted in

a 96 well microplate. In this case, when replacing the

waste tray, the 96 well microplate has to be placed on

the empty blue collection tube rack (Figure 4b).

• For increased DNA concentration, decrease the elution

volume to 60 μl, as per QIAGEN's recommendations, in

the VIALINK software.

Figure 4: Elution into a) provided collection

microtubes or b) a 96 well microplate.

A)

B)

CHAPTER 3: Application Notes 65

Remarks

Vacuum manifold:

Alignment of the vacuum manifold is very important in this process. Adding marks on the deck

helps to reposition the manifold whenever needed. To check the position of the well plate on top

of the vacuum manifold, attach the tips manually to the pipette. The pipette tips should be in the

middle of the wells. If not, adjust the position of the vacuum manifold on the deck.

Automatic mode:

The VIAFLO 96 can also operate in hands-free automatic mode, allowing the user to have

more walk-away time and less interaction, which is highly beneficial when using the instrument

in a laminar flow cabinet. The customized automatic VIALINK program can be found on the

INTEGRA website.

Conclusion

• The VIAFLO 96 electronic handheld pipette allows fast and simple liquid transfers for high

throughput PCR product purification.

• Optimized pipette settings enable accurate sample and reagent transfer, without the tip

touching and scratching the QIAquick membrane.

• The VIAFLO 96 electronic handheld pipette's compact design takes up minimal space

and fits on any lab bench.

• The unique operating concept makes the VIAFLO 96 and VIAFLO 384 as easy to use as

a conventional electronic pipette.

• The QIAvac 96 manifold is easily placed on the instrument and allows the processing of

other kits using 96 well silica-membrane or filter plates.

• Another option for this application is the MINI 96, which is the most affordable 96 channel

option on the market.

For more information

and a list of materials

used, please refer to

our website.

66 CHAPTER 3: Application Notes

3.5 PCR purification with Beckman Coulter

AMPure XP magnetic beads and the VIAFLO 96

Automatic magnetic bead purification with the VIAFLO 96

handheld electronic pipette

Agencourt AMPure XP magnetic beads (Beckman Coulter) are an efficient

way to clean up samples for PCR, NGS, cloning and microarrays. The kit

provides a solution for medium to high throughput requirements when carried

out in a 96 well plate, but the protocol involves many washing and transfer

steps that make it tedious to perform manually. With the VIAFLO 96,

a handheld 96 channel electronic pipette, multistep protocols such as

PCR clean-up and DNA purification can be

performed quickly and efficiently, increasing

throughput tremendously by transferring

samples and reagents to all 96 wells at once.

Thanks to its unique operating concept,

the VIAFLO 96 remains as easy to use as

a traditional handheld pipette and can even

provide critical information (user-defined

prompts) about the protocol steps.

Key benefits

• The VIAFLO 96 enables transfer of

samples, reagents and wash solutions to

96 wells at once, increasing the throughput

of magnetic bead-based DNA purification

methods.

• The partial tip loading of the VIAFLO 96

allows purification of fewer than 96 DNA

samples if necessary; 8, 16, 24, 32, 40

or 48 GRIPTIPS can be loaded for easy

purification of different numbers of samples.

• The optimal immersion depth for removing

supernatant or adding liquid right onto

the samples is guaranteed by defining the

z-height of the VIAFLO 96.

• The Tip Align setting of the VIAFLO 96

automatically positions the tips in the center

of the wells of a 96 well plate, avoiding any

disturbance of the beads.

CHAPTER 3: Application Notes 67

Overview: How to automate PCR purification steps

with VIAFLO 96

The VIAFLO 96 handheld electronic pipette with a three position stage is used to purify DNA

with AMPure XP beads from Beckman Coulter. The following protocol is an example of a set-up

for 96 samples, where each well of a 96 well plate is filled with 10 μl of DNA sample and 18 μl

of AMPure XP beads, then further processed with the VIAFLO 96. The PCR purification can

be performed manually or semi-automated using the VIAFLO 96 in automatic mode.

Custom-made VIALINK programs are provided. The VIALINK programs are set up according

to the manufacturer’s protocol (AMPure XP Beckman Coulter).

Step-by-step procedure

1. Dispense AMPure XP beads into PCR tubes

Transfer AMPure XP beads from the stock solution into 12 PCR tubes placed in

a cooling block from INTEGRA.

Note: The cooling block is just used as a support in this instance, not for cooling down the

samples.

To ensure a homogenous stock solution, beads are thoroughly mixed by shaking/inverting until

the solution appears consistent in color. The beads are transferred into 12 PCR tubes using

the Repeat Dispense mode of a VIAFLO single channel 1250 μl electronic pipette. A customized

VIALINK program (AMP_Transfer1) is available to aid bead transfer.

For optimal pipetting, ensure beads are thoroughly mixed before each transfer. Mixing steps

can be defined by the number of cycles and the pipetting speed. Both influence the efficiency

of mixing and thus the quality of the

clean-up. Saving these parameters in the

pipetting program ensures that mixing is

always carried out as defined, yielding

consistent results. Insert a pre- and

post-dispense step to enhance accuracy

and precision while pipetting precious

reagents, such as AMPure XP beads.

Tip: The use of sterile, filter, low retention

GRIPTIPS ensures that every dispense

is as accurate as possible, with no loss of

beads or sample.

Figure 1: Transfer AMPure XP beads from the stock solution into 12 PCR

tubes.

68 CHAPTER 3: Application Notes

2. Transfer AMPure XP beads into the DNA samples

Transfer AMPure XP beads from the PCR tubes into a 96 well plate preloaded

with DNA samples.

Pipette the beads from the PCR tubes

into the 96 well plate using a VIAFLO

12 channel 50 μl electronic pipette. For

optimal pipetting, make sure the tips are

exchanged, and mix the beads thoroughly

before each transfer. A customized

VIALINK program (AMP_Transfer2) is

provided for this step.

Tip: Use low retention GRIPTIPS to

minimize loss of beads adhering to the

tip wall.

3. Mixing and binding of the AMPure XP beads

Mixing and binding of the magnetic beads to the PCR samples.

Load GRIPTIPS (position A) then select

and run the AMPure_XP_M program on

the VIAFLO 96. The samples are now

mixed 10 times by pipetting up and down

on position B. A five minute wait time

follows, timed by the VIAFLO 96, to allow

the DNA to bind to the beads.

Tip: Use the z-height setting of the

VIAFLO 96 to define the optimal tip

immersion depth. This prevents air

entering the tip during mixing and avoids

the pipette tip touching the bottom of the

plate. Setting the Tip Align support strength to 3 for positions A and B makes it more comfortable

to use the VIAFLO 96. These settings can be incorporated into the program so that they are not

forgotten.

Figure 2: Transfer AMPure XP beads from the PCR tubes into a 96 well

plate preloaded with DNA samples.

Figure 3: Mixing and binding of the magnetic beads to the PCR samples.

CHAPTER 3: Application Notes 69

4. Magnetic separation of the AMPure XP beads

Separating the magnetic beads from the PCR samples.

Note: Make sure new GRIPTIPS are loaded

before continuing the protocol to ensure

removal of the supernatant without bead

carryover.

A prompt on the pipette screen reminds the

user to move the sample plate from position

AB onto the 96 well magnet (position B) and

place an automation friendly reagent reservoir

for waste collection on position AB. After a two

minute incubation time, the beads form a ringshaped

structure and the solution becomes

clear. Load new GRIPTIPS before continuing the procedure to ensure accurate removal of

the supernatant without bead carryover. Follow the instructions on the pipette and aspirate

the supernatant slowly from the sample, dispensing it into the waste reagent reservoir

(position AB).

Tip: To avoid disturbing the ring of beads, the supernatant is aspirated slowly at speed 1.

Leave 5 μl of supernatant in the plate to prevent beads being drawn out during aspiration.

The z-height limit is again used to ensure that the beads are not disturbed during pipetting.

5. AMPure XP bead clean-up

Wash the magnetic beads twice with 70 % ethanol.

Place an automation friendly reagent reservoir

containing 70 % ethanol on position A and

change the GRIPTIPS before continuing

with the wash step. Follow the prompts on

the pipette. Pre-wet the GRIPTIPS with 70 %

ethanol. Then wash the samples with 70 %

ethanol. Repeat the washing step again as

indicated by the pipette.

Tip: Pre-wetting the GRIPTIPS with 70 %

ethanol ensures equilibration of the humidity

and the temperature between the air in the

pipette/tips and the sample/liquid. In-house testing has shown that low retention GRIPTIPS

prevent ethanol from dripping while traveling from one pipetting position to another.

Figure 4: Separating the magnetic beads from the PCR samples.

Figure 5: Wash the magnetic beads twice with 70 % ethanol.

70

6. Elute samples from the magnetic beads

Elute the purified samples from the magnetic beads by adding the elution

buffer.

As indicated by the pipette, replace the 70 %

ethanol reagent reservoir on position A with

an elution buffer reagent reservoir and move

the sample plate from the magnet (position B)

to position AB. Load new GRIPTIPS before

continuing with the protocol. After transferring

and thoroughly mixing the elution buffer with

the beads, the pipette prompts the user to

place the sample plate back onto the magnet

(position B). During the one minute incubation

time, place a new 96 well plate on position AB.

7. Transfer the sample eluates

Transfer the sample eluates into the new 96 well plate.

Note: Load new GRIPTIPS to ensure a clean

eluate transfer without bead carryover.

Continue with the same program, slowly and

carefully transferring the eluates from position

B into the new plate (position AB).

Tip: Optimizing pipette settings (aspiration

speed, volume and height) allows the volume

of the transferred eluate to be maximized

without carryover of beads. These settings

can be easily tweaked at any time. Performing

a test run with water before implementing any

new assay is an ideal way to optimize pipette settings.

Figure 6: Elute samples from the magnetic beads.

Figure 7: Transfer the sample eluates into the new 96 well plate.

CHAPTER 3: Application Notes

CHAPTER 3: Application Notes 71

Remarks

Automatic mode:

The VIAFLO 96 can also operate on its own,

enabling less user interaction, which in turn

improves ergonomics and reproducibility. This

also makes it even more ideal for use in tight

spaces, such as under a laminar flow cabinet.

Partial tip load:

If you are not working with a full set of 96

samples, the VIAFLO 96 is able to work with

any number of tips loaded, allowing purification

of smaller numbers of samples. Figure 8: Automatic mode and partial tip load.

Conclusion

• The VIAFLO 96 is perfectly suited to magnetic bead purification in a 96 well format. An

entire plate with 96 samples can be purified in a fraction of the time it would take with a

traditional pipette.

• Optimized tip immersion and pipette settings in combination with the use of low retention

GRIPTIPS allow maximum sample recovery at the end of the purification protocol.

• The VIAFLO 96 can guide the user through the entire protocol step by step, ensuring the

correct workflow and enhancing the reproducibility of results.

• The optional automatic mode of the VIAFLO 96 enables the instrument to operate on its

own to minimize pipetting errors, making it even more ideal for use under a laminar flow

cabinet.

For more information

and a list of materials

used, please refer to

our website.

72 CHAPTER 3: Application Notes

3.6 PCR purification with Beckman Coulter

AMPure XP magnetic beads and

the ASSIST PLUS

Automatic magnetic bead purification with

ASSIST PLUS pipetting robot

Agencourt AMPure XP beads (Beckman Coulter) are used

for DNA purification in a variety of applications, including PCR,

NGS, cloning and microarrays. The ASSIST PLUS pipetting

robot provides a solution for optimal bead

separation and maximized recovery of

precious samples. User guidance

throughout the entire protocol

ensures an error-free pipetting

procedure. Careful and accurate

handling of the magnetic beads

by the ASSIST PLUS leads to

superior reproducibility and consistency

during the experiment. Taken together, the

ASSIST PLUS provides researchers with an easy

and highly efficient way to purify DNA from PCR reactions using AMPure XP magnetic beads.

Key benefits

• The VIAFLO and VOYAGER electronic

pipettes, in combination with

ASSIST PLUS, provide unmatched

pipetting ergonomics.

• Optimal pipette settings, including tip

immersion depth, pipetting speeds and

angles, maximize reproducibility and

sample recovery.

• Exact positioning of the pipette tips in the

sample wells avoids the risk of disturbing

the ring of magnetic beads or bead

carryover.

• The ASSIST PLUS automates many steps

of a magnetic bead purification protocol

and guides the user through the remaining

manual operations to ensure an error-free

process.

CHAPTER 3: Application Notes 73

Overview: How to automate PCR purification steps

with ASSIST PLUS

The ASSIST PLUS is used to purify DNA samples using AMPure XP beads (Beckman Coulter).

The pipetting robot runs a VOYAGER 8 channel 125 μl electronic pipette with 125 μl sterile,

filter, low retention GRIPTIPS. The use of low retention GRIPTIPS guarantees optimal liquid

handling of viscous (AMPure XP buffer) and volatile (70 % ethanol) solutions.

Below is an example set-up for 24 samples, preparing 10 μl DNA samples (position B) with

18 μl of AMPure XP beads (position A). The pipetting programs were prepared according to the

manufacturer’s protocol (AMPure XP, Beckman Coulter) using VIALAB software.

The protocol is divided into two programs that guide the user through every step of the PCR

purification process.

• Program 1: Binding (AMP_BINDING)

• Program 2: Washing and elution (AMP_WASH_ELUTE)

Experimental set-up: Program 1

Deck position A: PCR 8 tube strip containing the AMPure XP

beads (Figure 1, blue), placed onto a cooling

block from INTEGRA. Note: the cooling block

is just used as a support in this instance, and

not for cooling down the samples.

Deck position B: 96 well plate with 24 DNA samples for

purification (Figure 1, green).

Deck position C: 96 well ring magnet.

74 CHAPTER 3: Application Notes

Figure 1: Pipetting schema, set-up for program 1.

A B C

Run program 1: transfer & binding

Select and run the AMP_BINDING program on the VOYAGER electronic pipette. The

ASSIST PLUS pipetting robot immediately starts the protocol.

1. AMPure XP transfer

Transferring AMPure XP beads from an 8 tube PCR strip to a 96 well plate

containing the DNA samples.

To ensure the AMPure XP buffer is homogenous, the beads are resuspended by pipetting up

and down 10 times before being transferred to the samples. The beads and DNA fragments

are thoroughly mixed together before the pipette automatically starts the timer for a 5 minute

incubation, ensuring optimal conditions for the DNA strands to bind onto the magnetic beads.

Tip: Using low retention GRIPTIPS rather than regular GRIPTIPS prevents the loss of AMPure

XP beads during the pipetting steps (see Figure 2).

VOYAGER 8 channel

125 μl

50/125 μl sterile, filter,

low retention GRIPTIPS

PCR 8-Tube Strip on cooling

plate – 200 μl

96 well plate Sapphire

– 200 μl

96 well plate Sapphire on 96 well ring

magnet – 200 μl

CHAPTER 3: Application Notes 75

Figure 2: The image highlights the advantages of using low retention GRIPTIPS versus regular

GRIPTIPS when pipetting AMPure XP beads.

Figure 3: The beads and DNA fragments are thoroughly mixed together before the incubation.

76 CHAPTER 3: Application Notes

2. Magnetic separation of the AMPure XP beads

Separating the magnetic beads from the PCR samples.

A message instructs the user to move the plate (position B) onto the magnet (position C).

Continue the program to start the timer. After a two minute incubation on the magnet the

beads form a ring in the sample well and the solution becomes clear. The program resumes

automatically, and the supernatant is removed. On completion of this step, the pipette prompts

the user to continue with the AMP_WASH_ELUTE program and to replace the labware on

position A with the 8 row polypropylene (PP) reagent reservoir containing the ethanol and

elution buffer.

Tip: The supernatant is aspirated slowly using the Tip Travel feature of the ASSIST PLUS

to avoid disturbing the ring of beads. The Tip Travel feature keeps the tip immersion depth

constant during aspiration and dispensing. 5 μl of supernatant remain in the plate to prevent

beads being drawn out during aspiration.

Figure 4: The ASSIST PLUS settings allow removal of the supernatant without any bead carryover.

CHAPTER 3: Application Notes 77

Experimental set-up: Program 2

Deck position A: The 96 well PCR cooling block is replaced by

an 8 row polypropylene (PP) reagent reservoir

filled with 70 % ethanol in row 1 (blue) and

elution buffer in row 2 (orange). Row 8 is used

for waste (purple).

Deck position B: Emtpy 96 well plate.

Deck position C: 96 well ring magnet and 96 well plate with 24

DNA samples for purification (green).

Figure 5: Pipetting schema, set-up for program 2.

VOYAGER 8 channel

125 μl

50/125 μl sterile, filter,

low retention GRIPTIPS 8 row reagent reservoir 96 well plate Sapphire

– 200 μl

96 well plate Sapphire on 96 well ring

magnet – 200 μl

A B C

78 CHAPTER 3: Application Notes

Run program 2: Washing & elution

Start the AMP_WASH_ELUTE program on the VOYAGER electronic pipette. The

ASSIST PLUS washes the beads twice by automatically adding and removing ethanol.

3. Magnetic bead clean-up

Washing the magnetic beads twice with 70 % ethanol.

The programmed pipette settings allow the beads to be washed without disturbing the bead

ring. At the end of the second washing step, all the ethanol is removed. If necessary, an

additional drying time can easily be added using VIALAB software.

Tip: The use of low retention GRIPTIPS prevents ethanol from dripping while traveling from

position A to position C (see Figure 6).

Figure 6: The image highlights the advantages of using low retention GRIPTIPS (left) versus regular

GRIPTIPS (right) when pipetting ethanol.

4. Elute samples from the magnetic beads

Eluting the samples from the magnetic beads by adding an elution buffer.

The pipette prompts the user to move the reaction plate from the magnet (position C) to position

B. Continuing the protocol, the ASSIST PLUS transfers the elution buffer to the DNA samples

bound to the magnetic beads (position B, orange). After mixing carefully and thoroughly 10

times, the pipette prompts the user to place the 96 well plate on the magnet (position C).

CHAPTER 3: Application Notes 79

5. Transfer the sample eluates

Transferring the sample eluates into a new 96 well plate.

As indicated by the pipette, place a new 96 well plate onto position B and continue the program.

The sample eluates are then transferred into the new plate automatically.

Tip: Optimized pipette settings (aspiration speed, volume, height, tip travel and tip touch) allow

the volume of eluate transferred to be maximized without carryover of beads (see Figure 6).

A tip touch after the transfer removes droplets that may still cling to the end of the pipette tips.

Pipetting heights on the ASSIST PLUS can be fine-tuned at any time. Performing a test run with

water before implementing any new assay is an ideal way to optimize pipette settings.

Results

Figure 7: Magnetic beads are clearly visible in the 96 well plate with no supernatant remaining.

80 CHAPTER 3: Application Notes

Figure 8: No carryover of beads is observed in the eluate.

Conclusion

• Magnetic bead purifications can be easily automated on the ASSIST PLUS pipetting

robot.

• Optimized tip immersion and pipette settings together with the use of low retention

GRIPTIPS allow maximum sample recovery at the end of the purification protocol.

• The pipette loaded onto the ASSIST PLUS prompts the user when needed, eliminating

the risk of human errors.

• VIALAB programs can be easily adapted to specific labware.

• Prolonged pipetting tasks lead to repetitive strain injury. This can be avoided by

automating these steps with the ASSIST PLUS.

For more information

and a list of materials

used, please refer to

our website.

CHAPTER 4: Customer Testimonials 81

CHAPTER 4:

Customer testimonials

Our range of innovative liquid handling products has helped countless laboratories to achieve

PCR success, improve their throughput and further their ground-breaking research. But don’t

just take our word for it! Here are a few stories from our satisfied customers, demonstrating why

INTEGRA Biosciences is the right choice for PCR pipetting solutions and labware.

4.1 INTEGRA pipettes – the obvious choice for

start-up PCR labs

The gradual reopening of the world following the pandemic has led to an unprecedented

demand for COVID-19 testing, with schools, universities and workplaces relying on negative

PCR tests to continue operating. Matrix Diagnostics – a dedicated COVID-19 testing lab in

California – is helping to fulfill this critical need, relying on INTEGRA’s EVOLVE and MINI 96

pipettes to streamline and accelerate PCR workflows.

PCR-based diagnostic testing is a well-established technique in clinical labs around the world,

and this method has been brought to the attention of every household as the gold standard for

COVID-19 testing. However, the public is less aware that the sensitivity of this technique makes

it time-consuming and troublesome to perform without the right tools, as it is very sensitive to

pipetting errors and cross-contamination.

Founded in January 2021, Matrix Diagnostics

was established to meet the growing demand

for PCR testing in the San Francisco Bay Area,

and the newly formed team understood the need

for effective pipetting solutions from the outset.

Fady Ettnas, Lab Manager at Matrix Diagnostics,

explained: “We realized that, to meet the

anticipated demand for testing, we would have to

turnover between 2000 and 5000 samples every

day. This seemed like an impossible task for a new

lab with limited resources but, after implementing

INTEGRA’s pipettes in our lab, we quickly

alleviated the pipetting bottlenecks, putting us on

track to achieve our targets.”

Photo courtesy of Matrix Diagnostics

82

Evolving workflows

“Our protocols involve a range of repetitive

pipetting steps – including mixing reagents

and serial dilutions – for thousands of

samples a day, which has the potential

to be a cumbersome and error-prone

task,” Fady continued. “We therefore

chose INTEGRA’s EVOLVE manual

pipettes and MINI 96 portable electronic

pipettes to improve the reproducibility

and productivity of our workflows. We

have a number of single channel EVOLVE

pipettes, covering volumes ranging from

0.2 to 5000 μl, as well as 8, 12, and 16

channel models. What I like most about

EVOLVE is its ergonomic design and

ability to set volumes in a flash. The

unique design of INTEGRA’s GRIPTIPS also means that they never leak or fall off, avoiding

cross-contamination and maintaining sterility. We also use the compact MINI 96 extensively,

which is especially well suited to PCR set-up. It saves a lot of time and effort – around 15

minutes per cycle – when performing the wash steps. And because we run more than 25 cycles

every day, this is a huge saving, allowing us to process a much higher number of samples. It is

a perfect and affordable solution for our needs.”

A long-term investment

The benefits of these pipettes to users, particularly in terms of preventing physical strain

caused by repeated pipetting actions, are a priceless advantage. “I think the pipettes are a

great investment with huge returns, allowing the team to process more samples and improving

their pipetting experience. The company’s customer service is quick, responsive and helpful

and, crucially, the team was able to advise us on the right choice of pipettes to meet our

workload and objectives.”

Planning future with INTEGRA

“Currently, we are only offering COVID-19 tests, but we plan to expand to include other tests

including sexually transmitted diseases, urinary tract infections and flu, and we know that we

will need to automate our workflow. We will need something flexible and incredibly efficient and,

therefore, we are planning to acquire an ASSIST PLUS pipetting robot. I like all the INTEGRA

products that I’ve used, and have rarely encountered even minor technical issues. I think they

are the most obvious pipetting choice for both for start-ups and established lab set-ups, and are

well worth the investment,” Fady concluded.

Photo courtesy of Matrix Diagnostics

CHAPTER 4: Customer Testimonials

83

Photo courtesy of Harvard Medical School

4.2 A better qPCR pipetting experience

Manual pipetting can be a major bottleneck for research laboratories, especially when they face

the challenge of combining accurate results with high throughput. Like all repetitive tasks that

require precise actions, filling multiwell plates by hand is time consuming, and physically and

mentally draining, which can lead to errors. When Daisy Shu joined the Saint-Geniez laboratory

at Harvard Medical School, her experience was quite different, thanks to the INTEGRA VIAFLO

electronic pipettes.

From patients to pipettes

After graduating in optometry from the University of New South

Wales in Sydney, Daisy worked as an optometrist for two years

before deciding to pursue a PhD in cataract research at the

University of Sydney. She explained: “The move from my usual

clinical work with patients to research was a big change for me,

as I had to dive deep into molecular biology. I didn't even know

how to use a pipette back then! Cataracts – clouding of the

eye’s lens – are a leading cause of blindness worldwide, and I

studied their formation and ways to prevent that happening. My

focus was on transforming growth factor beta (TGF-β), which

has an important role in cancer metastasis, but is also relevant

for certain types of cataracts. I looked at the different signaling

pathways it activates and how those pathways interlink.”

Daisy completed her PhD in January 2019, and straight

afterwards flew to Boston to work as postdoctoral fellow in the

Saint-Geniez laboratory, continuing her research into eye health.

Here, she was able to apply her knowledge of TGF-β to agerelated

macular degeneration (AMD). Daisy continued: “I'm now

looking at how TGF-β causes the retinal mitochondria to change morphology and become

dysfunctional, altering cellular metabolism. The research is still at an early stage, so we're

mainly trying to understand how to prevent AMD, but the end goal is to find a cure.”

A better pipetting experience

At Harvard, Daisy was introduced to VIAFLO electronic pipettes, which were a complete

contrast to the large, fully automated pipetting workstation she had used during her PhD

research. The laboratory was already using two VIAFLO pipettes – a 125 μl eight channel

pipette and a 12.5 μl single channel version – and their flexibility compared to the automated

workstation dramatically improved her pipetting experience. “Complete automation on a large

workstation has its place, but there are downsides,” said Daisy. “You have to program every

CHAPTER 4: Customer Testimonials

84

single step perfectly before you can click one button and run the

protocol, and the process of fine-tuning takes a long time.”

“I found the VIAFLO pipettes amazing. A lot of our work is PCRbased,

performed in 384 well plates, and the VIAFLO pipettes

are real lifesavers. I use the 8 channel VIAFLO for most qPCR

liquid transfers, and the single channel pipette to add the

primers. Once you've made your master mixes and programmed

the pipette, it's really fast; it only takes me 20 minutes to

do a complete 384 well plate. When I was using the robotic

workstation in Sydney, I used to think that doing a qPCR was

really a big deal. Now, with the INTEGRA pipettes, it's just

so easy.”

VIAFLO pipettes provide a choice of pipetting modes and allow

easy adjustment of parameters such as volume and speed, as

well as providing pre-set programs and the option for custom

workflows. This helps laboratories to reduce errors and increase

throughput and reproducibility regardless of the users’ pipetting

experience. For Daisy, VIAFLO electronic pipettes have become the standard for how pipetting

should be: “In any pipetting workflow, you have to get every step right first time, otherwise you’d

end up having to troubleshoot the assay and do it again. I'm really surprised when I hear people

from other labs say they pipette each well individually with manual single channel pipettes. I’m

sure that would take forever compared to electronic pipetting, and my eyes would really suffer.

The VIAFLOs make everything easy. I love the color coding – it makes it so simple to match the

right tip to the right pipette – and the instrument can even be set to alert you when you need to

pipette again.”

CHAPTER 4: Customer Testimonials

Photo courtesy of Harvard Medical School

85

4.3 COVID-19 – Accelerate your PCR set-up

The emergence and outbreak of the novel coronavirus SARS-CoV-2 (COVID-19) has placed

unprecedented demands on laboratories testing patient samples for COVID-19, leaving

scientific staff to contend with a spiraling influx of COVID-19 samples and a rapid, continuous

growth in workload. Among the challenges faced by the Microbiology and Molecular Pathology

Department at Sullivan Nicolaides Pathology (SNP) – part of the Sonic Healthcare Group

– in Brisbane, Australia, is the increased pressure on laboratory automation used for both

coronavirus and pre-existing respiratory virus panel testing.

As a result of the coronavirus pandemic, SNP found itself analyzing extreme numbers of

samples, which exhausted the capacity of its automation platforms. At the same time, staff

were faced with a need to spend more time working up new virus testing protocols, which

were often performed manually or using semi-automated methods to fast track test response

times, leaving them prone to increased ergonomic strain. There was a clear need for additional

automated liquid handling instruments to increase sample processing capacity, reduce manual

intervention by laboratory analysts and fast track assay development for COVID-19 sample

testing.

Working together

In early March 2020, Kelly Magin and James Sundholm from

INTEGRA’s Australian distributor, BioTools Pty Ltd, partnered

with Shane Byrne, Scientific Department Head, Microbiology and

Molecular Pathology Department, SNP, to support COVID-19 testing

of patient samples using the ASSIST PLUS pipetting robot. An

ASSIST PLUS automated pipetting protocol was developed and

validated, enabling samples to be prepared in low volume, 384 well

plates for subsequent processing on a rapid, high throughput,

plate-based, real-time PCR amplification and detection instrument.

A VOYAGER adjustable tip spacing pipette and low retention

GRIPTIPS were used to transfer one-step RT-PCR master mix from a

low dead volume (<20 μl) SureFlo 10 ml reagent reservoir into a 384

well plate. The VOYAGER pipette also allowed automatic transfer

and reformatting of nucleic acid template extracted from combined

nasopharyngeal/oropharyngeal flocked swab(s) or sputum samples,

from 4 x FluidX™ 1.0 ml 96 format tube racks into the 384 well plate. The total PCR reaction

volume was reduced to 10 μl; 7.5 μl one-step RT-PCR master mix and 2.5 μl of nucleic acid

template. This miniaturization doubled the available testing capacity and simultaneously

reduced consumption of expensive one-step RT-PCR reagents of dwindling availability, with

associated cost savings.

Photo courtesy of Sullivan Nicolaides

Pathology

CHAPTER 4: Customer Testimonials

86

Defining success

SNP successfully validated the automated

protocol against its existing manual

processing method, performed using a

handheld electronic pipette. The results

were shown to be reproducible, precise

and accurate, with no contamination

observed in either the control or patient

samples. The compact, easy-to-use

ASSIST PLUS pipetting robot, complete

with validated protocol, was fully deployed

within five working days. While the current

protocol uses 384 well plates, it can be

readily adapted to 96 well format to meet

future needs.

4.4 Reducing protocol time for PCR using

96 channel pipette

Implementing an INTEGRA VIAFLO 96 electronic pipette has enabled the Virus- and Prion

Validation (VPV) Department at Octapharma Biopharmaceuticals GmbH, (Frankfurt, Germany)

to reduce the time taken to undertake PCR assays by greater than 60 %.

Since its foundation in 1983, Octapharma has been committed to patient care and medical

innovation. Its core business is the development and production of human proteins from human

plasma and human cell-lines.

The VPV Department has been set-up to investigate pathogen inactivation and removal steps

along the manufacturing processes. Among other techniques, multi-step 96 well format PCR

assays were developed, which involve three washing steps twice in the protocol. To undertake

their PCR assay more efficiently, Octapharma sought a system that enabled reproducible and

accurate liquid handling in the 96 well format and was able to completely remove residual liquid

as well as avoid well-to-well contamination.

Dr. Andreas Volk, a research scientist at Octapharma Biopharmaceuticals commented: "The

classical liquid handling solutions, fully automated robots or ELISA plate washers were either

too costly or prone to cross contamination in a PCR assay." He added: "When we tested the

INTEGRA VIAFLO 96 channel pipette, it fully met our requirements as it enabled mediumthroughput

liquid handling while minimizing cross-contamination. Additionally, the

Photo courtesy of Sullivan Nicolaides Pathology

CHAPTER 4: Customer Testimonials

87

VIAFLO 96 electronic pipette provided all the

adjustment options, which we had been used

to with manual pipettes, plus a specified tip

immersion depth for each pipetting step. With

our PCR protocol, which involves ten full liquid

transfers per plate, we now only use half the

amount of pipette tips as we can use the same

tips for liquid addition and aspiration in each

washing step. VPV Department staff has found

using the VIAFLO 96 benchtop pipette highly

intuitive and the overall time required for our

PCR washing procedures has been reduced to

approximately one third of the original time."

The INTEGRA VIAFLO 96 is a handheld 96

channel electronic pipette that has struck a

chord with scientists looking for fast, precise

and easy simultaneous transfer of

96 samples from microplates without the

cost of a fully automated system. The

VIAFLO 96 was designed to be handled just

like a standard handheld pipette – a fact

borne out by consistent end user feedback

that no special skills or training are required to

operate it. Users immediately benefit from the

increased productivity delivered by their VIAFLO 96. Fast replication or reformatting of 96 and

384 well plates and high precision transferring of reagents, compounds and solutions to or from

microplates with the VIAFLO 96 is as easy as pipetting with a standard electronic pipette into

a single tube. Four pipetting heads with pipetting volumes up to 12.5 μl, 125 μl, 300 μl or

1250 μl are available for the VIAFLO 96. These pipetting heads are interchangeable within

seconds enabling optimal matching of the available volume range to the application performed.

For 384 well pipetting, an enhanced version is available with VIAFLO 384. It features

384 channel pipetting heads in the volume range of 12.5 μl and 125 μl and is compatible with

96 channel pipetting heads.

Dr. Andreas Volk, Octapharma Biopharmaceuticals

CHAPTER 4: Customer Testimonials

88

CHAPTER 5:

Conclusion

So, there you have it, a full run down of PCR. By now, you should have all the information you

need to become a PCR pro, but if you’d still like to learn more about this interesting topic, we

have a wealth of articles on our website. Whatever your PCR requirements, we at INTEGRA

Biosciences are always available to answer your questions and provide you with the best

workflow solutions.

CHAPTER 5: Conclusion

89

CHAPTER 6:

References

1.1 The complete guide to PCR

1. Crow, E. (2012). Mind Your P's And Q's: A Short Primer On Proofreading Polymerases.

https://bitesizebio.com/8080/mind-your-ps-and-qs-a-short-primer-on-proofreadingpolymerases

2. Kim, S. W. et al. (2008). Crystal structure of Pfu, the high fidelity DNA polymerase from

Pyrococcus furiosus. International Journal of Biological Macromolecules, 42(4), 356-

361. https://doi.org/10.1016/j.ijbiomac.2008.01.010

3. ThermoFisher Scientific (n.d.). PCR Setup – Six Critical Components to Consider.

https://www.thermofisher.com/ch/en/home/life-science/cloning/cloning-learningcenter/

invitrogen-school-of-molecular-biology/pcr-education/pcr-reagents-enzymes/

pcr-component-considerations.html

4. AAT Bioquest (2020). What is the function of MgCl2 in PCR?

https://www.aatbio.com/resources/faq-frequently-asked-questions/What-is-thefunction-

of-MgCl2-in-PCR

5. Lorenz, T. C. (2012). Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting

and Optimization Strategies. Journal of Visualized Experiments, 63, e3998.

https://doi.org/10.3791/3998

6. Merck (n.d.). Polyermase Chain Reaction.

https://www.sigmaaldrich.com/CH/en/technical-documents/technical-article/

genomics/pcr/polymerase-chain-reaction

7. Viana, R. V., Wallis, C. L. (2011). Good Clinical Laboratory Practice (GCLP) for

Molecular Based Tests Used in Diagnostic Laboratories. In Akyar, I. (Ed.), Wide

Spectra of Quality Control (29-52). InTech.

https://cdn.intechopen.com/pdfs/23728/InTech-Good_clinical_laboratory_

practice_%20gclp_for_molecular_based_tests_used_in_diagnostic_laboratories.pdf

8. Ogene M. (2021). How does ddPCR work?

https://mogene.com/how-does-ddpcr-work

9. ThermoFisher Scientific (2016). Real-time PCR handbook.

https://www.ffclrp.usp.br/divulgacao/emu/real_time/manuais/Apostila%20qPCRHandbook.

pdf

CHAPTER 6: References

90

10. Prediger, E. (2017). Digital PCR (dPCR) – What is it and why use it?

https://eu.idtdna.com/pages/technology/qpcr-and-pcr/digital-pcr

11. Bio-Rad Laboratories (n.d.). Introduction to Digital PCR.

https://www.bio-rad.com/en-uk/life-science/learning-center/introduction-to-digital-pcr

12. Bio-Rad Laboratories (n.d.). Digital PCR and Real-Time PCR (qPCR) Choices for

Different Applications.

https://www.bio-rad.com/en-uk/life-science/learning-center/digital-pcr-and-real-timepcr-

qpcr-choices-for-different-applications

13. Schoenbrunner, N. J. et al. (2017). Covalent modification of primers improves PCR

amplification specificity and yield. Biology Methods and Protocols, 2(1).

https://doi.org/10.1016/j.ijbiomac.2008.01.010

14. Merck (n.d.). Hot Start PCR.

https://www.sigmaaldrich.com/CH/en/technical-documents/technical-article/

genomics/pcr/hot-start-pcr

15. Parichha, A. (2021). Nested PCR || Principle and usage.

https://www.youtube.com/watch?v=nHCjgo2Ze0o

16. New England Biolabs (n.d.). FAQ: What is touchdown PCR?

https://international.neb.com/faqs/0001/01/01/what-is-touchdown-pcr

17. Parichha, A. (2021). Touch down PCR.

https://www.youtube.com/watch?v=s9oV2-53esA

18. Cheriyedath, S. (2018). History of Polymerase Chain Reaction (PCR).

https://www.news-medical.net/life-sciences/History-of-Polymerase-Chain-Reaction-

(PCR).aspx

19. Arney, K. (2020). The Story of PCR.

https://geneticsunzipped.com/news/2020/11/3/the-story-of-pcr

20. Biosearch Technologies (2022). Taq facts.

https://blog.biosearchtech.com/thebiosearchtechblog/bid/48174/taq-facts

21. National Museum of American History (n.d.). Mr. Cycle, Thermal Cycler.

https://americanhistory.si.edu/collections/search/object/nmah_1000862

CHAPTER 6: References

91

1.2 Simple PCR tips that can make or break your success

1. Cheriyedath, S. (2018). History of Polymerase Chain Reaction (PCR).

https://www.news-medical.net/life-sciences/History-of-Polymerase-Chain-Reaction-

(PCR).aspx

2. Seeding Labs (2019). How To: PCR Calculations.

https://www.youtube.com/watch?v=CnQV5_CEvAo

3. McCauley, B. (2020). Setting Up PCR Reactions.

https://brianmccauley.net/bio-6b/6b-lab/polymerase-chain-reaction/pcr-setup

4. New England Biolabs (n.d.). Guidelines for PCR Optimization with Taq DNA

Polymerase.

https://international.neb.com/tools-and-resources/usage-guidelines/guidelines-forpcr-

optimization-with-taq-dna-polymerase

5. Lorenz, T. C. (2012). Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting

and Optimization Strategies. Journal of Visualized Experiments, 63, e3998.

https://doi.org/10.3791/3998

6. Gold Biotechnology (2020). How To: PCR Master Mixes.

https://www.youtube.com/watch?v=LSfvCJ9gUQU

1.3 Setting up a PCR lab from scratch

1. Bustin, S. A., Benes, V., Garson, J. A., et al (2009). The MIQE Guidelines: Minimum

Information for Publication of Quantitative Real-Time PCR Experiments. Clinical

Chemistry, 55(4), 611–622.

https://doi.org/10.1373/clinchem.2008.112797

2. National Human Genome Research Institute (2020). Polymerase Chain Reaction

(PCR) Fact Sheet.

https://www.genome.gov/about-genomics/fact-sheets/Polymerase-Chain-Reaction-

Fact-Sheet

3. Viana, R. V., Wallis, C. L. (2011). Good Clinical Laboratory Practice (GCLP) for

Molecular Based Tests Used in Diagnostic Laboratories.

https://cdn.intechopen.com/pdfs/23728/InTech-Good_clinical_laboratory_practice_

gclp_for_molecular_based_tests_used_in_diagnostic_laboratories.pdf

4. Redig, J. (2014). The Devil is in the Details: How to Setup a PCR Laboratory.

https://bitesizebio.com/19880/the-devil-is-in-the-details-how-to-setup-a-pcrlaboratory

5. Mifflin, T. E. (n. d.). Setting Up a PCR Laboratory.

https://pubmed.ncbi.nlm.nih.gov/21357132/

CHAPTER 6: References

92

6. Gu, M. (n. d.). Molecular Laboratory Design And Its Contamination Safeguards.

https://www.scimmit.com/molecular-laboratory-design-and-its-contaminationsafeguards

7. Lee, R. (2015). Molecular Laboratory Design, QA/QC Considerations.

https://www.aphl.org/programs/newborn_screening/Documents/2015_Molecular-

Workshop/Molecular-Laboratory-Design-QAQC-Considerations.pdf

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work

1. Bustin, S. A., Benes, V., Garson, J. A. et al. (2009). The MIQE guidelines: minimum

information for publication of quantitative real-time PCR experiments. Clinical

Chemistry, 55(4), 611-622.

https://doi.org/10.1373/clinchem.2008.112797

2. Rutledge, R. G., Côté, C. (2003). Mathematics of quantitative kinetic PCR and the

application of standard curves. Nucleic Acids Research, 31(16).

https://www.gene-quantification.de/rudledge-2003.pdf

3. Applied biological materials (2016). Polymerase chain reaction (PCR) – Quantitative

PCR (qPCR).

https://www.youtube.com/watch?v=YhXj5Yy4ksQ

4. Nagy, A., Vitásková, E., Černíková, L. et al. (2017). Evaluation of TaqMan qPCR

system integrating two identically labelled hydrolysis probes in single assay. Scientific

reports, 7.

https://doi.org/10.1038/srep41392

5. Bradburn, S. (n.d.). How to calculate PCR primer efficiencies.

https://toptipbio.com/calculate-primer-efficiencies

6. Bio-Rad (n.d.). qPCR assay design and optimization.

https://www.bio-rad.com/en-ch/applications-technologies/qpcr-assay-designoptimization?

ID=LUSO7RIVK

7. University of Western Australia (2016). Melt curve analysis in qPCR experiments.

https://www.youtube.com/watch?v=FvJnXKzejSQ

8. Bio-Rad (2011). Real time QPCR data analysis tutorial.

https://www.youtube.com/watch?v=GQOnX1-SUrI

9. Bio-Rad (2011). Real time QPCR data analysis tutorial (part 2).

https://www.youtube.com/watch?v=tgp4bbnj-ng

10. Kannan, S. (2021). 4 easy steps to analyze your qPCR data using double delta Ct

analysis.

https://bitesizebio.com/24894/4-easy-steps-to-analyze-your-qpcr-data-using-doubledelta-

ct-analysis

CHAPTER 6: References

93

1.5 How to design primers for PCR

1. Benchling (n.d.). Primer Design.

https://www.benchling.com/primers

2. Addgene (n.d.). How to Design a Primer.

https://www.addgene.org/protocols/primer-design

3. PREMIER Biosoft (n.d.). PCR Primer Design Guidelines.

http://www.premierbiosoft.com/tech_notes/PCR_Primer_Design.html

4. Merck (n.d.). Oligonucleotide Melting Temperature.

https://www.sigmaaldrich.com/CH/en/technical-documents/protocol/genomics/pcr/

oligos-melting-temp

5. Integrated DNA Technologies (n.d.). How do you calculate the annealing temperature

for PCR?

https://eu.idtdna.com/pages/support/faqs/how-do-you-calculate-the-annealingtemperature-

for-pcr

CHAPTER 6: References

INTEGRA Biosciences AG

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HOW TO BECOME A PCR PRO

The polymerase chain reaction (PCR) is a key life sciences technique. It has been used in

molecular biology – including molecular diagnostics – for many years, and a number of different

types, for example, RT-PCR, qPCR, vPCR and ddPCR have been developed over time.

Today, PCR is a vital tool for the detection of pathogens, such as the SARS-CoV-2 virus, and

is essential for genotyping and NGS library preparation. However, PCR is well known for being

difficult to run successfully and several parameters must be considered when planning the PCR

protocol.

We have therefore compiled this eBook – consisting of in-depth educational articles, relevant

app notes and customer testimonials – to help you understand how PCR works, and what needs

to be considered to perform effective PCR reactions. We also demonstrate how our solutions

can help you to enhance the throughput of your lab, and become a PCR pro in no time.

Dr Éva Mészáros

Application Specialist

eva.meszaros@integra-biosciences.com

Anina Werner

Content Manager

anina.werner@integra-biosciences.com

FOREWORD

TABLE OF CONTENTS

CHAPTER 1: What you need to know about PCR

1.1 The complete guide to PCR 2

1.2 Simple PCR tips that can make or break your success 15

1.3 Setting up a PCR lab from scratch 20

1.4 qPCR: How SYBR® Green and TaqMan® real-time PCR assays work 24

1.5 How to design primers for PCR 32

CHAPTER 2: INTEGRA Biosciences’ PCR solutions 38

CHAPTER 3: Application notes

3.1 Efficient and automated 384 well qPCR set-up with the ASSIST PLUS pipetting robot 42

3.2 Automated RT-PCR set-up for COVID-19 testing 49

3.3 Increase your sample screening and genotyping assay throughput with the VOYAGER 57

adjustable tip spacing pipette

3.4 PCR product purification with QIAquick® 96 PCR Purification Kit and the 61

VIAFLO 96 handheld electronic pipette

3.5 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 66

VIAFLO 96

3.6 PCR purification with Beckman Coulter AMPure XP magnetic beads and the 72

ASSIST PLUS

CHAPTER 4: Customer testimonials

4.1 INTEGRA pipettes – the obvious choice for start-up PCR labs 81

4.2 A better qPCR pipetting experience 83

4.3 COVID-19 – Accelerate your PCR set-up 85

4.4 Reducing protocol time for PCR using 96 channel pipette 86

CHAPTER 5: Conclusion 88

CHAPTER 6: References 89

2

CHAPTER 1:

What you need to know about PCR

In this chapter, we will cover topics such as PCR’s fascinating history, its mechanism and

different variations, and techniques for troubleshooting common issues you may encounter.

We’ll also go through tips for establishing a PCR lab, as well as a comprehensive overview of all

things related to qPCR and primer design.

1.1 The complete guide to PCR

Polymerase chain reaction (PCR) methods have been carried out in labs around the world since

the 1980s, opening the door for an array of new applications, such as genetic engineering,

genotyping and sequencing. In this article, we take a deep dive into this fascinating technique

by explaining its mechanism, exploring its history, looking into the different types of PCR,

discussing troubleshooting tips and much more.

CHAPTER 1: What you need to know about PCR

3

What is PCR?

The polymerase chain reaction (PCR) is a fast and inexpensive technique for amplifying a DNA

sequence of interest. It consists of three steps:

• Denaturation: The sample is heated to separate the DNA into two single strands.

• Annealing: The temperature is lowered to allow primers to anneal to specific single-stranded

DNA segments, flanking the sequence to be amplified.

• Extension: The temperature is raised to the optimum working temperature of the polymerase

enzyme, which then makes a complementary copy of the DNA sequence of interest.

One such repetition or 'thermal cycle' theoretically doubles the amount of the DNA sequence of

interest present in the reaction. Typically, 25 to 40 cycles are performed – resulting in millions

or even billions of DNA copies – depending largely on the amount of DNA in the starting sample

and the number of amplicon copies needed for post-PCR applications.

The three steps of a PCR reaction are carried out automatically by a thermal cycler, but can

only be successful if the master mix has been correctly prepared. The following sections

explain the components that make up the master mix and how they interact with the template

DNA during thermal cycling.

PCR master mix components

The PCR master mix consists of six components:

• PCR-grade water: Certified to be free of contaminants, nucleases and inhibitors.

• dNTPs: Containing equal concentrations of the four nucleotides (dATP, dCTP, dGTP and

dTTP), which are the 'building blocks' to create complementary copies of the DNA sequence

of interest.

• Forward and reverse primers: Short, single-stranded DNA sequences that anneal

specifically to the plus and minus strands of the template DNA, flanking the sequence to

be amplified. For some assays – such as protocols amplifying much-studied genes or DNA

sequences of common bacteria – ready-to-use primers can be purchased. However, many

experiments require custom PCR primers tailored to the region of interest of the template DNA

and the reaction conditions.

CHAPTER 1: What you need to know about PCR

4

• DNA polymerase: Taq-polymerase is the most commonly used enzyme for PCR reactions.

It uses dNTPs to create complementary copies of the DNA sequence of interest. For some

applications, such as mutagenesis, Taq-polymerase is not accurate enough and the use

of high fidelity polymerases is recommended. Just like Taq-polymerase, they sometimes

add an incorrect nucleotide when replicating the template DNA but, as they have a 3' to

5' exonuclease activity, they 'proofread' the newly synthesized strands and correct any

mistakes.1 This proofreading step is highly beneficial for accuracy but it also slows down PCR

reactions, and high fidelity polymerases (also called slow polymerases) therefore need about

twice the time of Taq-polymerase to create a complementary DNA strand. The most popular

high fidelity DNA polymerase is Pfu-polymerase.2

• Buffer: Provides a suitable environment for the DNA polymerase, with a pH between 8.0

and 9.5.3

• Magnesium chloride: Increases the activity of the DNA polymerase and helps primers

to anneal to the template DNA for a higher amplification rate.4 This cofactor is sometimes

included in the buffer in a sufficient concentration.5

The template DNA , which may be genomic DNA (gDNA), complementary DNA (cDNA) or

plasmid DNA (pDNA), is then added after master mix preparation.

The 3 steps of PCR

After preparing the PCR master mix and adding the template DNA samples to it, you can load

your reaction tubes, PCR strips or microplates into the thermal cycler. They will then go through

the following steps:

• Denaturation: The thermal cycler first heats the reaction mix to 95-98 °C to denature the

template DNA, separating it into two single strands. Depending on your sample, this usually

takes 2-5 minutes during the first thermal cycle, and 10-60 seconds for subsequent cycles.

• Annealing: When the temperature is lowered, the primers anneal to the sequences flanking

the template DNA region of interest. Depending on the sequence and melting temperature of

your primers, this step usually takes 30-60 seconds, and the optimal annealing temperature

typically lies between 45 and 60 °C.

• Extension: The temperature is increased to 72 °C, which is the ideal working temperature

for the Taq-polymerase. Depending on the synthesis rate of your polymerase, and the length

of the target sequence, it usually takes 20-60 seconds to create complementary copies

of the DNA sequence of interest.6 After approximately 25-40 cycles – depending on the

amount of DNA present at the start, and the number of amplicon copies needed for post-PCR

applications7 – the last extension step should be extended to 5 minutes or longer, allowing the

Taq-polymerase to finish the synthesis of uncompleted amplicons.5 If you can't immediately

take your samples out of the thermal cycler after the final extension step because you're busy

with other experiments, program it to cool your samples to 4 °C. For overnight runs where you

CHAPTER 1: What you need to know about PCR

5

leave your samples in the thermal cycler for hours after the final extension step, you should

opt for a holding temperature of 10 °C instead of 4 °C, as it causes less wear and tear on your

machine.

As shown in the image above, the amount of PCR product theoretically doubles at every

thermal cycle, leading to an exponential increase of PCR product. However, in reality, the phase

of exponential amplification eventually levels off and reaches a plateau because the reagents

have been consumed and the DNA polymerase activity decreases.

The different types of PCR

After performing a standard PCR reaction, you can determine the concentration, yield and

purity of the amplified DNA sequences using gel electrophoresis, spectrophotometry or

fluorometry. However, you can’t determine the amount of template DNA present in a sample

before amplification using standard PCR. If this is a requirement for your experiment, you have

to perform a qPCR reaction.

qPCR

qPCR – also called real-time PCR, quantitative PCR or quantitative real-time PCR – is a

technique used to detect and measure the amplification of target DNA as it is produced.

In contrast to conventional PCR reactions, qPCR requires a fluorescent intercalating dye

or fluorescently-labeled probes, and a thermal cycler that can measure fluorescence and

calculate the cycle threshold (Ct) value. Typically, the fluorescence intensity increases

proportionately to the concentration of the PCR product being formed, measuring quantities

of the target in real time.

CHAPTER 1: What you need to know about PCR

6

qPCR can be divided into dye-based methods (e.g. SYBR® Green) and probe-based methods

(e.g. TaqMan®).

RT-PCR and RT-qPCR

Another limitation of standard PCR is that it can only be used to amplify DNA sequences. If you

want to amplify RNA target sequences, you have to use RT-PCR.

RT-PCR

vPCR

Reverse transcription PCR (RT-PCR) is used to amplify RNA target sequences, such as

messenger RNA or RNA virus genomes. This type of PCR involves an initial incubation of

the RNA samples with a reverse transcriptase enzyme and a DNA primer – comprising

sequence-specific oligo (dT)s or random hexamers – prior to the PCR amplification.

For viability PCR (vPCR), each sample needs to be split into two aliquots. One aliquot is

incubated with a photoreactive intercalating dye that is unable to diffuse through intact cell

membranes of live cells. This means that it only intercalates into the DNA of dead cells. When

this aliquot is subsequently treated with a blue light, the dye binds irreversibly to the DNA. Both

aliquots are then subject to DNA purification and qPCR amplification. If they exhibit similar

qPCR signals, the target microorganisms in the sample are mostly viable. If the dye-treated

aliquot exhibits a weaker signal, the target microorganisms are mostly dead. vPCR is an

important technique in diagnostics, agriculture and food safety.

You can also perform a qPCR reaction instead of executing a standard PCR reaction after

the reverse transcription step, which produces cDNA from RNA. This PCR variant is called

RT-qPCR.

vPCR

The third limitation of standard PCR is that it cannot distinguish between the DNA of viable

and non-viable cells. You should use vPCR if this is important to your application, for example,

because you want to know if the infectious microorganisms in a clinical sample are dead or

alive.

CHAPTER 1: What you need to know about PCR

7

ddPCR

Digital droplet PCR (ddPCR) is another relatively new type of PCR. It uses fluorescently labeled

probes to detect DNA sequences of interest, and a water-oil emulsion system to split each

sample into about 20,000 nanoliter-sized droplets. After amplification, every droplet of the

sample is analyzed on its own. Droplets that contain at least one DNA sequence of interest emit

a fluorescent signal – and are consequently positive – while droplets without the DNA sequence

of interest don't fluoresce, and are therefore negative. Using the Poisson distribution, you can

then determine the concentration of the DNA sequence of interest in the original sample by

analyzing the ratio of positive to negative droplets for absolute quantification.8

An advantage of ddPCR compared to qPCR is that it's more precise. While qPCR can detect

two-fold differences in DNA target sequence variation, e.g. discriminate 1 copy from 2 copies

of a gene, ddPCR can discriminate 7 copies from 8 copies, which means that it can detect

differences as small as 1.2-fold.9 On top of that, ddPCR is better suited for multiplexing assays

if you want to determine the ratio of low abundance to high abundance DNA sequences of

interest, such as rare mutations on wild type backgrounds. When using qPCR, the fluorescent

signal from the high abundance sequences can dominate and obscure the signal from the

low abundance sequences. This risk is ruled out with ddPCR, as each droplet behaves as

an individual PCR reaction and contains either zero, one or, at most, a few sequences of

interest.10,11

ddPCR

CHAPTER 1: What you need to know about PCR

8

Due to these advantages, ddPCR is often preferred over qPCR for the detection of mutations

and SNPs (single nucleotide polymorphisms), allelic discrimination, gene expression studies,

and the analysis of copy number variations.12

Hot start PCR

If your PCR reaction results in non-specific amplification, you can try to increase the reaction

specificity using a hot start polymerase. This enzyme remains inactive during master mix

preparation and sample addition at room temperature, eliminating the risk that unintended

products and primer dimers are formed during PCR set-up.13

Nested and semi-nested PCR

Nested or semi-nested PCR are alternatives to hot start PCR that increase reaction specificity.

Nested PCR uses two sets of primers and two successive PCR reactions. The first set of

primers is designed to amplify a DNA sequence slightly longer than the sequence of interest.

During the second PCR reaction, the second set of primers that is specific to the sequence of

interest anneals to the amplicons of the first PCR reaction and helps to amplify the sequence of

interest.14,15

Nested PCR

CHAPTER 1: What you need to know about PCR

9

Semi Nested PCR

Semi-nested PCR works similarly to nested PCR. During the first PCR reaction, one primer

anneals to the sequence of interest and the second primer to a region flanking the sequence of

interest. This primer is then replaced with a second primer annealing to the region of interest

during the second PCR reaction.

The idea behind nested and semi-nested PCR is that, if non-specific products were amplified

during the first PCR reaction, these will not be amplified during the second PCR reaction, as the

primers cannot anneal to them.

Touchdown PCR

A third type of PCR developed to increase reaction specificity is touchdown PCR. The assay

set-up for touchdown PCR is identical to the set-up for standard PCR. The only difference lies in

the annealing step. During the first thermal cycle, the annealing temperature should be several

degrees above the optimal primer annealing temperature, then be lowered by 1-2 °C every

second cycle.16 These high temperatures during the first cycles avoid PCR primers forming

primer-dimers or binding to regions outside the DNA sequence of interest. The downside is

that the PCR primers don't all sufficiently bind to the template DNA, which leads to low yields.17

However, this can be tolerated, as the low yield of specific amplicons is then exponentially

amplified with every thermal cycle that is performed at the optimal annealing temperature.

CHAPTER 1: What you need to know about PCR

10

The history of PCR

As we've shown, there are many different types of PCR, and some of them have only recently

been developed. However, the foundation for PCR was laid in the 1950s:

• In 1953, James Watson and Francis Crick discovered the double-helix structure of DNA, and

suggested that there might be a possible copying mechanism for DNA.

• Four years later, Arthur Kornberg identified the first DNA polymerase that was able to copy the

template DNA, although only in one direction.

• In 1971, Gobind Khorana and his team started to work on DNA repair synthesis. Their

technique used DNA polymerase repeatedly, but employed only a single primer template

complex, which did not allow exponential amplification.

• At the same time, Kjell Kleppe from Khorana's lab proposed a two primer system that would

double the amount of DNA in a sample, but no one actually conducted the experiment to

find out whether it worked. The reason for this was probably that there was not yet a DNA

polymerase that could withstand the high temperatures of the denaturation step. This means

that they would have had to add a fresh dose of enzyme after every thermal cycle.

• In 1983, Kary Mullis, working at Cetus Corporation, added a second primer to the opposite

strand, and realized that repeated use of DNA polymerase triggers a chain reaction that will

amplify a specific DNA sequence, thus inventing PCR. The patent got approved in 1987, and

he won the Nobel Prize in Chemistry six years later.

• In 1976, the thermostable enzyme Taq-polymerase – which is typically used in PCR today

– was first isolated from the bacterium Thermus aquaticus, which had been discovered in a

hot spring of Yellowstone National Park in 1969. When it was finally incorporated into PCR

workflows in 1988, it removed the need to add a new dose of enzyme after every thermal

cycle, paving the way for the invention of automated thermal cyclers.18,19,20

CHAPTER 1: What you need to know about PCR

11

PCR troubleshooting

One of the most important troubleshooting mechanisms is to always include positive and

negative control samples.

If the sequence of interest wasn't amplified in your positive control sample, your master mix,

template DNA or thermal cycler could be the source of the problem:

• Master mix: Have you added the right volume and concentration of each reagent, and have

you cooled your reagents during master mix preparation?

• Template DNA: Have you run an agarose gel to ensure that your template DNA isn't

degraded? Is your template DNA pure enough and, if not, have you purified it?

• Thermal cycler: Is the number of thermal cycles sufficient for your assay? Have you

programmed the device correctly, and is it calibrated to ensure that it performs the reaction

steps at the right temperatures?

If the sequence of interest was amplified in your negative control sample, one or more

components of your master mix is contaminated. PCR reactions are very sensitive, and create

large number of copies of DNA sequences from minute amounts of starting material, so

contamination is a common issue. To prevent it, the right lab set-up is crucial.

CHAPTER 1: What you need to know about PCR

12

Lab set-up

Ideally, your PCR lab should have two rooms, each divided into two areas. The first room should

be exclusively used for pre-PCR activities, and divided into a master mix preparation area and a

sample preparation area. The second room should have a dedicated area for amplification, and

another one for product analysis.

If you’re lacking in space or budget for a two-room PCR lab, you can set up the pre-PCR and

amplification and analysis areas in the same room, but ensure they are as far from one another

as possible. In addition to the spatial separation, you could also consider setting up your PCR

reactions in the morning, and performing the amplification and analysis steps in the afternoon.

Temporally separating the different steps of your PCR reactions may limit your flexibility and

make you lose some time, but lowers the risk of aerosols with high DNA concentrations from the

analysis area contaminating your master mix and samples in the pre-PCR area.

On top of these precautionary measures, you should always work in biosafety cabinets or

laminar flow hoods when setting up your PCR reactions, use different sets of pipettes for master

mix preparation, sample preparation and analysis, and make sure that you use filter tips and

consumables that are free of DNase, RNase and PCR inhibitors.

Specificity

Another major PCR challenge is specificity. As explained before, it can be improved using hot

start, nested, semi-nested or touchdown PCR. A further option to prevent the amplification of

regions outside the DNA sequence of interest, as well as the formation of secondary structures,

is to redesign your primers.

CHAPTER 1: What you need to know about PCR

13

Use this checklist to see whether your primers meet all the requirements:

• Are your primers between 18 and 24 bp long?

• Is your target sequence length between 100 and 3000 bp for standard PCR assays, or 75

and 150 bp for qPCR assays?

• Do your primers have melting temperatures between 50 and 60 °C, and within 5 °C of

each other?

• Have you performed a gradient PCR to determine the optimal annealing temperature?

• Does the GC content of your primers lie between 40 and 60 %?

• Have you avoided runs or repeats of four or more bases or dinucleotides?