Q5/Phusion PCR

Reaction Setup

1. Assemble all reaction components on ice/cold block in 250 µL "PCR" tubes (unless using Q5 Hot-Start Polymerase which doesn't require keeping cold).
   1. Enzyme must be added after at least buffer and water are mixed.
   2. Make master mixes when possible, as it reduces pipetting steps, reduces errors from pipetting small volumes, and maximizes component precision across reactions. Combine all common components for n reactions together and into reaction tubes aliquot reaction volumes less the volume of the variable component.
      e.g., primers templates commonly vary across reactions, so for ten 25 µL reactions containing a combined volume of 0.75 µL primers + template, combine water, buffer, dNTPs, any enhancers, and polymerase; mix and aliquot 24.25 µL of this master mix across ten tubes, before adding their unique primers and templates. Components may be multichannel-pipetted into reactions for convenience.
2. After adding last component, mix reaction with pipette or by closing, flicking, and centrifuging tubes to recollect liquid at bottom.
3. Transfer the reactions to a thermocycler preheated to the denaturation temperature (98°C). Pausing just after starting the program will pause the protocol after the lid has finished (slowly) heating, after which pausing will pause the program with the block heated to the denaturation temp, step 1.

Reaction Guidelines

Template

Use of high quality, purified DNA templates greatly enhances the success of PCR. Recommended amounts of DNA template are:

Low quality genomic DNA preparation

Low quality genomic DNA is present in cell lysate and is often sufficient for cloning PCR or strain genotyping. Simply "boil" a cell solution in a thermocycler tube for 10 min, centrifuge the cell debris, and use the supernatant as genomic DNA. The cell solution can be a saturated liquid culture or a mass of cells resuspended in TE (elution buffer). This lysate can be frozen and reused many times as a PCR template.

Primers

Oligonucleotide primers are generally 17–40 nucleotides in length and ideally have a GC content of 40–60%. Computer programs such as Primer3 (in Benchling) can be used to design or analyze primers. The best results typically come from reactions with 0.5 µM each primer.

For small reactions, the 100 µM primer volume can be <0.2 µL and thus not accurately pipettable. Using a minimum of 0.2 µL, though excess, is typically still successful and convenient as it avoids making primer dilutions. Primer dilutions often waste time and freezer space, given that they aren't typically needed once the immediate PCR is complete.

Buffers

The 5× Q5 Reaction Buffer provided with the enzyme is recommended as the first-choice buffer for robust, high-fidelity amplification.

The 5× Q5 Reaction Buffer is detergent-free and contains the optimal 2.0 mM MgCl₂ at the final (1×) concentration. Shyam deduced based on safety data sheets that Q5 Reaction Buffer, but not Phusion buffer, contains glycerol, which reduces DNA secondary structure, and tetramethylammonium, which increases primer stringency.

Store buffers at 20°C for long-term. Once you claim an aliquot, you may store it at 0–4°C until it is used up, preventing the need for thawing.

Deoxynucleotides

The final concentration of dNTPs is typically 200 μM of each deoxyribonucleotide, typically mixed and stored as a 10 mM solution at -20°C. Freeze-thaw cycles of dNTPs must be limited to preserve the triphosphate moiety, just as with ATP-containing solutions. dNTP stocks are thus aliquoted in 0.25 or 0.6 mL tubes in 20 µL volumes.
Q5 High-Fidelity DNA Polymerase cannot incorporate dUTP and is not recommended for use with uracil-containing primers or templates.

Q5 DNA Polymerase concentration

Q5 High-Fidelity DNA Polymerase is recommended to be used at a final concentration of 20 U/mL (1.0 U/50 μl reaction). However, the optimal concentration of Q5 High-Fidelity DNA Polymerase may vary from 10–40 U/mL (0.5–2 U/50 μl reaction) depending on amplicon length and difficulty. Do not exceed 2 U/50 μl reaction, especially for amplicons longer than 5 kb.

The hot-start formulation, Q5 Hot-Start DNA Polymerase, inhibits the robust exonuclease (and polymerase?) activity of the enzyme, allowing for convenient room temperature reaction setup. The aptamer/inhibitor is released from the enzyme during normal cycling conditions, so no separate activation step is required.

PCR Additives

Difficult PCRs with GC-rich sequences or secondary structure in the amplified DNA or primers may be improved by the addition of Q5 High GC Enhancer at a final 10–20% (from the 5–10× stock). It is not a reaction buffer itself and cannot be used alone, only to be added to reactions along with reaction buffer when other conditions have failed. NEB says use of the Q5 High GC Enhancer often lowers the range of temperatures at which specific amplification can be observed, but that the normal, unmodified T_anl generally still supports specific amplification. It is unclear how most PCR additives below influence polymerase fidelity.

A [Mg²⁺] of 2.0 mM is optimal for most PCR products generated with Q5 polymerase, which is provided by 1× Q5 Reaction Buffer and should not require further supplementation.

Shyam deduced that Q5 reaction buffer contains glycerol, which reduces DNA secondary structure, and tetramethylammonium, which increases primer stringency; and Q5 High-GC Enhancer contains DMSO and glycerol to do more of the same.

PCR Additives:

• DMSO reduces DNA secondary structure. Use at 3–10% final, adjusting in 2% increments. 10% DMSO lowers T_m 5.5–6°C. DMSO reduces the activity of Taq polymerase, which may need to be increased with high DMSO.^{(1)(2)(4)(5)(6)}
• Glycerol reduces DNA secondary structure. Found in Q5 buffer. Use at 5–10% final.⁽²⁾⁽⁵⁾
• Formamide increases the stringency of primer annealing, resulting in less non-specific priming and increased amplification efficiency. Use at 1-10%.^{(1)(2)(4)(5)(6)}
• Tetramethylammonium chloride increases the stringency of primer annealing, resulting in less non-specific priming and increased amplification efficiency. Use at 10–100 mM final.⁽¹⁾⁽²⁾
• Triton X-100, Tween-20/-40 or NP-40 reduce DNA secondary structure, but can increase non-specific amplification. Use at 0.1–1% final.⁽¹⁾⁽²⁾ TWEEN can neutralize SDS left over from template DNA preparation that would inhibit the reaction. Use at 0.25–1% final.⁽¹⁾⁽²⁾
• Betaine / Betaine·H₂O greatly reduces the high T_m bias of G:C over A:T pairs, reversing the bias slightly at lower T_ms. Especially useful for GC-rich templates. Use at 1–3 M final. Can inhibit amplification of some templates. Don't use betaine·HCl.^{(1)(2)(4)(5)(6)} Use 1 M final.⁽⁸⁾
• 1,2-propanediol greatly improves success of GC-rich template amplification. Use at 0.8 M final.⁽³⁾⁽⁷⁾
• Ethylene glycol greatly improves success of GC-rich template amplification. Use at 1 M final.⁽³⁾
• Sulfolane (tetramethylene sulfone) can improve success of GC-rich template amplification. Use at 0.4 M final.⁽⁶⁾
• 7-deaza-2′-deoxyguanosine is a dGTP analogue especially useful for extremely GC-rich templates. Success is reported with up to 83% GC. Use a 1:3 ratio of dGTP:7-deaza-2′-deoxyguanosine.⁽²⁾
• BSA prevents reaction components adhering to the tube. Use at ≤0.8 mg/mL final.⁽²⁾⁽⁵⁾
• Trehalose ^(4)(5)(7) Use 0.4 M final.⁽⁸⁾
• Dithiothreitol (DTT) ⁽⁴⁾⁽⁵⁾
• 5× preCES-II: 4 M betaine, 10 mM DTT, 10% DMSO.⁽⁴⁾
• "Nagai universal mix": 1 mg/mL BSA, 10 mM DTT, 5% glycerol.⁽⁵⁾
• "Horáková universal mix": final 1 M 1,2-propanediol, 0.2 M betaine.⁽⁷⁾

Sources:  (1) Bitesize Bio, (2) Bitesize Bio, (3) Bitesize Bio / Zhang, (4) Ralser, (5) Nagai, (6) CSH, (7) Horáková, (8) Spiess

Thermocycling

Note: If you need to destroy the non-synthetic template (plasmid), you can add 1 µL DpnI per 50 µL completed PCR, and incubate 37˚C, 15–60 min. DpnI has full activity in Phusion buffer and 50% activity in Q5 buffer ^NEB. 

Thermocycling Guidelines

Denaturation

An initial denaturation of 30 seconds at 98°C is sufficient for most amplicons from pure DNA templates. Longer denaturation times can be used (up to 5 min) for templates that require it.

During thermocycling, the denaturation step should be kept to a minimum. Typically, a 5–10 second denaturation at 98°C is recommended for most templates.

Annealing

Optimal annealing temperatures for Q5 High-Fidelity DNA Polymerase tend to be higher than for other PCR polymerases. The NEB Tm Calculator should be used to determine the annealing temperature when using this enzyme. Typically, use a 10–30 second annealing step at 3°C above the T _m of the lower T _m primer. A temperature gradient across a strip of aliquotted reactions can also be used to optimize the annealing temperature for each primer pair.

2-step PCR

When primers with an annealing temperature ≥ 70°C are used, a 2-step thermocycling protocol (combining annealing and extension into one 72° step) is possible. 

Touchdown/Touch-up PCR

See Troubleshooting

Extension

The recommended extension temperature is 72°C. Extension times are generally 20–30 seconds per kb for complex, genomic samples, but can be reduced to 10 seconds per kb for simple templates (plasmid, E. coli, etc.) or complex templates < 1 kb. Extension time can be increased to 40 seconds per kb for cDNA or long, complex templates, if necessary.
When amplifying products > 6 kb, it is often helpful to increase the extension time to 40–50 seconds/kb.

A final extension of 2 minutes at 72°C is recommended.

Cycle number

Generally, 25–35 cycles yield sufficient product. For genomic amplicons, 30-35 cycles are recommended.

PCR product

The PCR products generated using Q5 High-Fidelity DNA Polymerase have blunt ends. If cloning is the next step, then blunt-end cloning is recommended. If T/A-cloning is preferred, the DNA should be purified prior to A-addition, as Q5 High-Fidelity DNA Polymerase will degrade any overhangs generated.

Troubleshooting/optimizing a PCR

• Recheck primer annealing temperatures and GC-content.

• Add 5% DMSO or high GC-enhancer or other additives. See Additives section. A lower, 3% DMSO can be added as a preventative measure, or if significant homodimerization or heterodimerization of primers is expected, a few DMSO concentrations can be tested (0/5% or 3/6% or 3/6/9%) with a 2–5° lower T_anl.

• To increase specificity (remove spurious products), use the touchdown thermocycling protocol. See below.

• To increase specificity (remove spurious products or obtain a missing product), use the touch-up thermocycling protocol. See below.

• To increase specificity (remove spurious products or obtain a missing product), try a range of annealing temperatures. See Annealing section.

• Remake your primer stocks.
• If strong spurious template, ensure reagents are not contaminated. E. coli genomic contamination is common in cheaply purified polymerase.
• If no products at all, ensure polymerase, buffer, and dNTPs are functional in a positive control reaction with known-to-work primers on a trusted template.

Touchdown PCR

To enhance amplification specificity, a touchdown thermocycling protocol can be used, which starts at a higher, stringent T _anl and ramps it down across successive cycles to a steady, permissive T _anl, ensuring high specificity of primer binding in initial products at the elevated T _m, which have a head start in amplification. If the desired band is not visible at all under standard protocol, touch-up PCR must be used. PMID 1861999 PMID 8679209 

A typical protocol consists of (after initial denaturation) 10 PCR cycles with the expected T _anl +5°C decrementing 0.5°C per cycle, followed by 25 PCR cycles with T _anl. The T _{anl, i} must be at most the extension temperature. The constant, T _{anl, f} ought to be at least the expected T _anl and can be set as high as the T _anl of the entire primer oligo pair for ensured sustained high stringency. The number of ramping cycles must equal the T _anl elevation divided by the T _anl decrement, and the number of constant T _anl cycles must be ≈25 cycles to ensure sufficient amplification after ramping.

n _{cycles, ramp} = (10–15 cycles) = (T _{anl, i} – T _anl) / ΔT 
ΔT = (T _{anl, i} – T _anl) / (10–15 cycles)

Touch-up PCR

While the touchdown thermocycling enhances amplification specificity by imposing a more stringent initial annealing temperature and ramping to a more permissive T _anl across cycles, touch-up thermocycling does the opposite, by starting at the permissive (expected) T _anl and ramping up across successive cycles to a steady, higher, stringent T _anl, ensuring high specificity of primer binding in later products which are selectively amplified for over successive cycles from non-specific amplicons at the initial permissive T _anl. If the desired band does not appear at the expected T _anl, the initial T _anl can be lower so as to better ensure the correct amplicon is part of the initial selection pool. PMID: 22468135

A typical protocol can consist of (after initial denaturation) 10–15 PCR cycles with the expected (or lower) T _anl,i, incrementing ≥1°C to a final T _anl,f at least 5–10°C higher and as high as that of the entire oligo pair, followed by the remainder of PCR cycles using a T _anl,con equal to T _anl,f. T _anl,f can (perhaps should) be increased 2–5°C for higher threshold stringency, followed by the constant T _anl,con cycles not having that additional increase for better product priming. If the desired T _anl,const is ≥70–72°C, the constant phase can eliminate the annealing step (2-step PCR). If the primers do not have a non-annealing 5′ end or if the entire oligo pair T _anl is not substantially higher than that of just the annealing regions, then the T _anl,i cannot be elevated much without the artificial higher threshold T _anl,f being used for sufficient selectivity. Longer primers can be used or add on non-annealing 5′ ends to select for correct product by providing a T_m advantage to the product.

The 10–15 incrementing T _anl cycles must equal the T _anl elevation divided by the T _anl increment, and the number of constant T _anl cycles should be 20–25 cycles.

n _{cycles, ramping} = (10–15 cycles) = (T _{anl, f} – T _anl,i)/ΔT 
ΔT = (T _{anl, f} – T _anl,i)/(10–15 cycles)

Another kind of touch-up protocol cycles the set of T _anl ramping cycles 4–5 times; it has no constant T _anl phase. 

Primer Design

A section on general primer design may be included or may never.

Benchling Primer3Plus Sequencing Primer Design

These parameter calculations give good results for the design of sequencing primers with similar spacing as to manually-designed primers, but with the added benefit of template specificity checking. It will generate pairs of divergent forward and reverse primers, but often you'll only need just one of each pair to cover the entire target span with reads optimally spaced.

* Below, this 850 nt value represents your idea of reliable good sequencing read (after the ≈25 nt "junk" lead between the primer 3′ end and beginning of good sequencing trace). This spacing can be adjusted to the sequencing read length you're comfortable with, but be sure to adjust it in all the other parameters it's used here, marked with an * . 

• Select the target sequencing span; right click on the sequence (not the features), and in the menu click Run Primer3. 
• Task: Sequencing
• T_m Parameters:
  • Algorithm:
    • SantaLucia 1999, if sequencing or using Taq polymerase for a colony PCR using these primers
    • Modified Breslauer 1986 , if using Q5 or Phusion for a colony PCR
  • Click Set to Primer3 Defaults: DNA: 50 nM; Na⁺/K⁺: 50 mM; Mg²⁺: 1.5 mM; dNTP: 0.6 mM
• Region:
  • Target: Start  x to End  y  spanning the desired sequencing interval.
    
    These target indices need adjustment if the 5′ and 3′ ends will be within the coverage of existing sequencing primers (e.g. AB17/AB18 in the vector/connectors). Instead of simply omitting ~800 bp of the ends, it works better to include them in the equal partitioning of . To do this, select the full target interval (including flanking Golden Gate sites, where applicable), and click Use Selection. Calculate #Results R and Spacing S as below. Add this S to the START index and subtract S from the END index:
    x = left boundary index + S
    y = right boundary index – S
    Before you generate primers, you can reduce #Results by 2, or leave it alone and see an optional primer pair between the last two sequencing spans. 

• Primer:
  These parameters can likely be adjusted to your liking without issue. 
  
  • GC%: min 30% – opt 50% – max 70%
  • T_m: min 53° – opt 56° – max 63°
  • Size: min 17 nt – opt 20 nt – max 25 nt
  • 3′ GC Clamp: 1

• Result Generation:
  • # Results: R = ⌈L ÷ 850*⌉.
    #Results R  = (target length L) ÷ (850* nt reliable read length), rounded up, not down. Target length L = y – x target sequence indices, or just look at the length of the selection.

• Sequencing:
  
  • Spacing:  S = L ÷ R
    Spacing S = target seq length L ÷ #Results R. Normally between 575–900 nt. This evenly distributes the number of ideal primer sites across the target length.
  • Interval: 40 nt.
    If you need both primers in a primer pair that the results give you and they are too close to use (3′ ends <50 bp apart, reads may not overlap), then after saving other selected primer pairs, rerun Primer3 with Interval set to 50 or 60 nt to spread apart that primer pair.
  • Lead: 0 nt.
  • Accuracy: 20 nt