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  • AAV Titration by qPCR Using SYBR Green Technology

AAV Titration by qPCR Using SYBR Green Technology

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Introduction

The AAV Titration protocol can be used to determine the number of genome-containing particles of an AAV prep using SYBR green technology.

This protocol was validated using an internal reference AAV of known titer, 100837-AAV1, and by measuring the titer of samples obtained from academic viral vector cores. Our titers were similar to those reported by these institutions.

This protocol is for a 96-well plate with 20 μL reaction volume.

Last Update: February 13, 2019
Estimate of time required: ~3 h

Workflow Timeline

Plate set-up:
2 h
qPCR run:
1.5 h
Data analysis:
30 min

Equipment

  • qPCR instrument
  • Heating plate
  • Pipettors
  • 1–10 µL single channel pipette
  • 20–200 µL single channel pipette
  • 200–1000 µL single channel pipette
  • 1–10 µL multichannel pipette
  • 2–50 µL multichannel pipette
  • 20–200 µL multichannel pipette

Reagents

  • Universal SYBR Master Mix 2X
  • Primer pair targeting AAV2 ITR (Aurnhammer et al., 2012)
    • fwd ITR primer, 5'-GGAACCCCTAGTGATGGAGTT
    • rev ITR primer, 5'-CGGCCTCAGTGAGCGA
  • ITR-containing plasmid for standard curve
  • RNase-free DNase
  • 10X DNase buffer
  • Nuclease-free water
  • Microcentrifuge tubes
  • 96-well optical plate
  • Pipette tips

General Considerations

  • Always run standards and samples in duplicate at least
  • If possible, include an AAV reference sample of known titer. The reference material should have a titer within 1-log of the expected titer of the samples being tested.
  • Always include a No Template Control (NTC), i.e master mix + water
  • Whenever possible use a multichannel pipet to minimize pipetting error and variability
  • Mix samples very well by pipetting back and forth multiple times at each step

Reagent Preparation

Master Mix: Count the number of samples (n) and prepare master mix for an additional 10 samples (n+10 – the additional amount will ensure that there is enough master mix for all samples). Each sample requires 15 μL of master mix.

Pro-Tips
  • Use a "Universal" SYBR master mix which contains a high-quality DNA polymerase and a blend of dTTP/dUTP to minimize carryover contamination. The master mix should also contain an internal passive reference (typically ROX dye), to normalize non-PCR–related fluorescence fluctuations and to minimize well-to-well variability that result from a variety of causes, such as pipetting error and sample evaporation.
  • Make the master mix after all the samples have been added to the qPCR plate. Start by adding water, then SYBR master mix, then the forward and reverse primers. Vortex briefly, immediately before use.
  • Use a reservoir and a multichannel pipette to dispense the master mix into the wells.
Reagent Amount for ONE Reaction Amount for 100 reactions (1 x 96 well plate)
Universal SYBR Master Mix 2X 10 μL 1,000 μL
100 μM Forward Primer 0.15 μL 15 μL
100 μM Reverse Primer 0.15 μL 15 μL
Nuclease Free Water 4.7 μL 470 μL

Procedure

  1. Prepare a plasmid stock of 2 x 109 molecules/μL to generate a standard curve:
    • One option is to use plasmid #59462 from Addgene. The values highlighted below in red were calculated using this plasmid, but will change if you use a different plasmid.

      Sample Calculation

      Size in bp of Addgene plasmid #59462:
      6208 bp
      Concentration of Addgene plasmid #59462:
      1.07 μg/μL
      Molecular Weight:
      6208 bp x 650 dalton/bp or (g/mole)/bp = 4.03 x 106 g/mole
      Moles/μl:
      1.07 μg/μL x 1 g/106 μg x1 / 4.03 x 106 g/mole = 2.65 x 10-13 moles/μL
      Molecules/μl:
      2.65 x 10-13 moles/μL x 6.022145 x 1023 molecules/mole = 1.59 x 1011 molecules/μL
    • To obtain a solution at 2 x 109 molecules/μL:
      • 1.59 x 1011 / 2 x 109 = 79.8X dilution 100 μL / 79.8 = 1.25 μL
      • Therefore we need to dilute 1.25 μL stock into 98.74 μL H20
    Pro-Tip
    • Once a validated standard curve is obtained, make a small aliquot of each standard (enough for 1 or 2 plates) and store at -20 °C. Once a standard is thawed do not freeze it again but store at 4 °C and use within 1 week.
    • Keep track of the Ct value for each standard over time. They should remain within 0.5 Ct of their initial value. If the Ct value of the standard starts to drift, it’s time to make a new one.
    • When developing the assay multiple plasmids containing ITR were tested. Plasmid #59462 is one plasmid that gave reliable and consistent results. Use the recommended plasmid, or test multiple plasmids to find a suitable one.
    • Some labs have reported better results when the plasmid is linearized.
  2. Treat the purified AAV samples with DNase I to eliminate any contaminating plasmid DNA carried over from the production process (DNase does not penetrate the virion).
    1. 5 μL sample + 39 μL H2O + 5 μL 10X DNase buffer + 1 μL DNase
    2. Gently mix sample (do not vortex)
    3. Incubate 30 min at 37 °C
    4. Transfer to ice
    ** Critical: do NOT treat your plasmid standard with DNase **
  3. Make 6 serial dilutions, in duplicate, of your standard curve plasmid (2 x 109 stock made in step #1):
    Volume of 2 x 109 stock or previous dilution (μL) Volume of nuclease-free water (uL) Molecules per μL
    10 90 2 x 108
    10 of 2 x 108 dilution 90 2 x 107
    10 of 2 x 107 dilution 90 2 x 106
    10 of 2 x 106 dilution 90 2 x 105
    10 of 2 x 105 dilution 90 2 x 104
    10 of 2 x 104 dilution 90 2 x 103
    Pro-Tip
    To help stabilize the standards add carrier DNA to a final concentration of 4 ug/mL to each standard dilution.
  4. Dilute DNase-treated and AAV reference samples according to the dilution scheme in the table below:
    Dilution Series Volume of sample (uL) Volume of nuclease free water (uL) Dilution factor Total dilution
    Dilution 1 (DNase step) 5 uL AAV stock 45 uL 10X 10X
    Dilution 2 5 uL Dil. 1 95 uL 20X 200X
    Dilution 3 20 uL Dil. 2 80 uL 5X 1000X
    Dilution 4 20 uL Dil. 3 80 uL 5X 5000X
    Dilution 5 20 uL Dil. 4 80 uL 5X 25000X
    Dilution 6 20 uL Dil. 5 80 uL 5X 125000X
    Dilution 7 20 uL Dil. 6 80 uL 5X 625000X
    Dilution 8 20 uL Dil. 7 80 uL 5X 3125000X

    Dilutions highlighted in green are the ones loaded onto the qPCR plate for most samples.

    • If sample is expected to have a titer <1 x 1012 GC/mL, use dilutions 3–6
    • If sample is expected to have a titer >3 x 1013 GC/mL, use dilutions 5–8

    Note: at Addgene, samples typically range from 1 x 1012 GC/mL to >2 x 1013 GC/mL and we use an internal reference virus that is 1 x 1013 GC/mL. We recommend always using a reference within 1-log of the expected titer.

    Pro-Tips
    • The quality of the sample dilution series is critical. Make sure to pipet each dilution up and down at least 10 times, and use at least half of the final volume (mix with >50 uL if your well contains 100 uL)
    • Use a multichannel pipette to load the standards and samples onto the qPCR plate
  5. Set up and load the 96-well plate:
    1. Load 5 μL of each standard in duplicate
    2. Load 5 μL of each sample in duplicate. Do not forget to include a no template control (NTC = master mix + water).
    3. Add 15 μL of Master Mix per well and mix well by pipetting back and forth at least 5 times.
    4. Seal plate with transparent film.
    5. Centrifuge at 3,000 rpm for 2 min to bring the sample to the bottom of the tube.
    6. Run the following protocol in your qPCR instrument using SYBR detection:
      • 98 °C 3 min / 98 °C 15 sec / 58 °C 30 sec / read plate/ repeat 39x from step 3 / melt curve

    Example of plate set-up:

    1 2 3 4 5 6 7 8
    A 1.00 x 109 1.00 x 108 1.00 x 107 1.00 x 106 1.00 x 105 1.00 x 104 empty NTC
    B 1.00 x 109 1.00 x 108 1.00 x 107 1.00 x 106 1.00 x 105 1.00 x 104 empty NTC
    C AAV reference Sample 3
    D
    E Sample 1 Sample 4
    F
    E Sample 2 Sample 5
    F
  6. Perform data analysis using the instrument’s software. Determine the physical titer of samples (viral genomes (vg)/mL) based on the standard curve and the sample dilutions.
    Pro-Tip
    Make sure that the qPCR is valid by checking to the following:
    • Standard curve: R2(coefficient of correlation) ~ 1.0, E (efficiency of PCR) ~100% (90%–110% range is acceptable)
    • Baseline removal: all samples will have some small amount of background signal that is most evident during initial PCR cycles. This background signal must be removed to accurately determine differences between samples.
    • Melt curve analysis: a single peak should be seen. The presence of a second peak at a temperature of ~70–75 °C usually indicates the presence of primer dimers which can increase background signal and alter the Ct values of your samples.
    • Quality of your standard curve: you should observe differences in Ct values that make sense for your dilutions (~3.3 difference Ct for a 10-fold dilution is appropriate).
    • Quality of duplicates: Exclude duplicates from analyses if there is more than a 0.5 Ct difference between them.

Sample Data

Standard Curve for AAV Titration by qPCR

Figure 1: Example of a valid 8-point standard curve.

qPCR of AAV Vector

Figure 2: Example of the amplification plots obtained from an AAV sample. Each curve represents a dilution.

References

Aurnhammer C, Haase M, Muether N, Hausl M, Rauschhuber C, Huber I, Nitschko H, Busch U, Sing A, Ehrhardt A, Baiker A. Universal real-time PCR for the detection and quantification of adeno-associated virus serotype 2-derived inverted terminal repeat sequences. Hum Gene Ther Methods. 2012 Feb;23(1):18-28. (Link opens in a new window)PMID: 22428977