DEVELOPMENT OF A METHODOLOGY FOR THE APPLICATION OF
SYNTHETIC DNA IN STREAM TRACER INJECTION EXPERIMENTS
(SUPPORTING INFORMATION)
Jan Willem Foppena*, Judith Seopaa, Noel Bakobiea, and Thom Bogaarda,b
5
a
UNESCO-IHE Institute for Water Education, P.O. Box 3015, 2601 DA Delft, The Netherlands; e-mail:
j.foppen@unesco-ihe.org
b Delft
University of Technology, Water Resources Section, Delft, The Netherlands
*Corresponding
author
10
Introduction
This text provides background information on the preliminary steps in the protocol,
e.g. determining the type of batch container and the optimal DNA sampling technique.
Then, a detailed account is given on how we determined DNA concentrations,
15
including inhibition, standard curve efficiency, and individual amplification curve
efficiencies. Finally, the way in which we determined confidence intervals of DNA
concentrations is described.
Materials and Methods
20
Preliminary steps in the development of the protocol
The initial steps in the development of the protocol were to determine the effect of
batch containers and water sample taking techniques on synthetic DNA
concentrations, and to select an adequate type of container and a suitable sample
taking technique. In our laboratory, we tested glass bottles (250 ml), polyethylene
25
(PE) bottles (250 ml), and polypropylene co-polymer (PPCO) bottles (250 ml;
Nalgene, Thermo Fisher Scientific). To each of the bottles, 100 ml of MilliQ water
1
and 450 μl of a concentrated solution of T22 and T23 was added. The bottles were
hand shaken and samples were taken immediately. Thereafter, the bottles were placed
on an Innova 2100 orbital shaker at 100 rpm, and samples were taken every 1-2 hour
for a total duration of 10 hour. The results (not shown) indicated that DNA mass
5
recovery from the PE bottles and PPCO bottles was statistically not significantly
different (see paragraph on 'Assessing confidence intervals of DNA tracer
concentrations' below), while mass recovery from the glass bottles significantly
differed from the input mass. From this we concluded that both PE and PPCO bottles
could be used as containers for our batch experiments. However, since PPCO bottles
10
have a known very low affinity for absorbing DNA particles (Gaillard and Strauss,
2000), we chose to carry out our batch experiments with PPCO bottles.
Next, in a PPCO bottle with 100 ml of MilliQ water and 450 μl of a concentrated
solution of T22 and T23, we tested a number of different sampling taking techniques:
1) sample collection in a 2.0 ml polypropylene syringe (BD Plastipak), 2) sample
15
collection in the same type of syringe, but sample disposal through a 0.22 µm
cellulose acetate syringe filter (Membrane Solutions, part # SFCA013022N), 3)
sample collection in the same type of syringe, but sample disposal through a 1.0 µm
glass fiber syringe filter (Membrane Solutions, part # SFGF013100N), 4) sample
collection in a 1 ml low adhesion polypropylene disposable gradient filtertip without
20
surface charge to assure no binding of charged molecules like DNA, and equipped
with a Self Sealing Non Collapsing (SSNC) filter tip (Bioplastics, Landgraaf, The
Netherlands) to avoid cross-contamination. In all 4 cases, storage of the sample was in
2.0 ml Eppendorf vials made of M-type polypropylene with less than 1% DNA
adhesion (Bioplastics, Landgraaf, The Netherlands). The results (data not shown)
25
indicated that glass fiber filters retained a statistically significant fraction of the
2
synthetic DNA mass (T22 or T23), while cellulose acetate filters, known to release
dissolved organic carbon into the sample (e.g. Khan and Subramania-Pillaib, 2007),
apparently increased the efficiency of the qPCR reaction giving rise to recovered
DNA masses (T22 or T23) well in excess of 100%. Best results were obtained when
5
using no filter, either with a syringe or a filtertip. We chose to use method 4, the
filtertips, because of their known low DNA binding characteristics.
Determining DNA tracer concentrations
The PCR reaction itself takes place in a carefully defined mixture of PCR buffer,
10
primers, probe, and sample, up to a total volume of 20-50 µl in a so called 'well',
which is then inserted into a qPCR apparatus. In our research, for each brook injection
experiment, a standard curve was determined, as we expected that the quality of the
brook water could have an effect on the performance of the enzymatic qPCR reaction,
thereby affecting the determination of DNA concentrations. In addition, prior to each
15
experiment, of all brooks a water sample was taken and transported in a coolbox to
our laboratory, where they were analysed for cations on an ICP-MS (Thermo Fisher
Scientific XSeries 2).
For each standard curve sample, the final mixture per qPCR well consisted of 2 µl
10X PCR buffer (Sigma-Aldrich product # P2317 buffer consisting of 100 mM
20
Trizma©-HCl, pH 8.3 at 25 °C, and 500 mM KCl) , 0.8 µl of PCR grade MgCl2
solution (Sigma-Aldrich product # M8787 with a concentration of 25 mM), 0.125 µl
of forward primer and of reverse primer both at a final concentration of 62.5 nM, 0.3
µl probe at a final concentration of 150 nM, 1 µl Taq polymerase (Genscript product #
E00007) corresponding to a final enzyme concentration of 0.5 U per PCR well, and
25
0.25 µl dATP (Genscript product # C01577) , 0.25 µl dGTP (product # C01578), 0.25
3
µl dCTP (product # C01579), and 0.25 µl dTTP (product # C01580) at a final
concentration of 0.2 mM each. To this mixture 4.0 µl of sample with known DNA
marker concentration, 4.0 µl of brook water and 6.65 µl of diethyl pyrocarbonate
(DEPC) treated PCR grade water was added up to a total PCR well volume of 20 µl.
5
Samples from injection experiments were prepared in a similar fashion. The only
difference was that instead of adding 4.0 µl of sample with known DNA
concentration, 4.0 µl of brook water and 6.65 µl of diethyl pyrocarbonate (DEPC)
treated PCR grade water, in this case 4.0 µl of sample from the injection experiment
(with unknown DNA concentration), and 10.65 µl of DEPC treated PCR grade water
10
was used (again adding up to a total PCR well volume of 20 µl). All samples were
subjected to an amplification protocol consisting of an initial denaturation step of 3
min at 95 °C followed by 40 cycles of denaturation and annealing/extension. The
denaturation step was 95 °C for 15 seconds and the annealing/extension step was at 60
°C for 1 minute.
15
Standard curves were prepared covering a dynamic range of 6 to 7 10-fold serial
dilutions corresponding to DNA marker concentrations ranging between 10-107 DNA
particles/PCR well. All standard curves were prepared in duplicate. In addition, the
standard curve data were used to determine the efficiency E [in %] of the PCR
reaction at various concentrations as:
20
1
𝐸 = (10− 𝑠 − 1) ∙ 100
(S1)
whereby 𝑠 was defined as the slope of the fitted linear curve between (known)
concentrations [expressed as the logarithm of the number of DNA marker particles
4
per µl in a PCR well] and threshold cycle. Furthermore, the standard curve was used
to assess the occurrence of inhibition of the polymerase enzyme due to the presence of
inhibiting compounds in the brooks we used for the injection experiments. These
inhibiting compounds may be humic substances and/or complex polysaccharides,
5
usually present in brook waters. Generally, when inhibition occurs, owing to the large
dynamic range of qPCR, samples can simply be diluted, usually between 10-100
times, in order to 'dilute away' inhibitors and to generate high quality amplification
profiles with adequate efficiency. To each standard curve, a so called no template
control (NTC) was added. The purpose of the NTC was to check for pollution during
10
the process of preparing the mixture for PCR analysis and to assess the importance of
random amplification and the formation of primer-dimers. Of course, for completely
controlled assays, random amplification should be absent, but since we worked with
environmental waters from which we never knew exactly the composition, some
random amplification occurred, and therefore, the lowest reliable detection threshold
15
cycle had to be determined.
In addition to determining the qPCR efficiency from the slope of the log-transformed
standard curve, we also determined the efficiency of each individual amplification
curve, 𝐸𝑖 , for each sample by using LinRegPCR (Ramakers et al., 2003; Ruijter et al.,
2009) according to:
20
𝑁 = 𝑁0 (𝐸𝑖 )𝐶
(S2)
where 𝑁 is the number of target molecules or amplicons at the end of cycle number 𝐶,
and 𝑁0 is the number of amplicons at the beginning of the reaction. In this equation,
25
the efficiency 𝐸𝑖 of the reaction is a number between 1 and 2. LinRegPCR determines
5
a baseline fluorescence and does a baseline subtraction. Then a so-called Window-ofLinearity is set and PCR efficiencies per sample are calculated from the slope of each
individual amplification curve. The reason for using LinRegPCR was that we wanted
to use the sample specific qPCR efficiency as an additional check to evaluate the
5
goodness of the design of the DNA markers, since a well designed DNA marker
should have similar qPCR efficiencies across the entire range of concentrations.
All pipetting actions were carried out with Gilson Pipetmans (P10, P20, P100, P200)
with the same disposable gradient filtertips as described above. Storage of all samples
was in 1.5 ml Eppendorf vials made of M-type polypropylene with less than 1% DNA
10
adhesion (Bioplastics, Landgraaf, The Netherlands). The preparation of qPCR mix (all
compounds as described above, except sample) was carried out in one DNA
workstation (Kisker L020-GC), while the preparation of the qPCR wells was carried
out in another -similar- DNA workstation, located adjacent to the first one.
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Assessing confidence intervals of DNA tracer concentrations
Carrying out a qPCR analysis requires a number of pipetting actions of volumes
ranging from 4-500 µl. The preparation of the qPCR mix (see above) requires the
pipetting of 8 different compounds, and the preparation of a standard curve requires
the dilution of the original, purchased vial of synthetic DNA by pipetting 10 µl into 90
20
µl of TE buffer (pH = 8.0), and the 13 times serial repetition of this process in order to
arrive at ultra-diluted DNA concentrations consisting of 50 or less DNA particles per
well. Then, 4 µl of sample plus 16 µl of PCR mix is finally pipetted into a PCR well.
So, for the preparation of the most dilute DNA marker concentration, a total of 26 (=
2 x each serial dilution) + 8 (preparation of qPCR mix) + 2 (adding qPCR mix and
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sample to a qPCR well) = 36 pipetting actions are required! Depending on the
6
accuracy and precision of the pipet and pipetting system used, this gives rise to errors
in the resulting DNA concentration, and it is important to quantify these errors in
order to determine realistic confidence intervals of the DNA marker concentrations.
We identified 2 types of errors: 1. Serial dilution errors, or, perhaps more accurately,
5
the propagation of the error in serial dilutions, and 2. Random errors due to
differences in performance of the qPCR apparatus and/or due to minor variations in
the preparation of the qPCR mix or in adding qPCR mix and sample to a qPCR well.
Serial dilution errors
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The serial dilution error was determined by following Hedges (2002). Briefly, the
serial dilution error consists of two parts: 1. Pipetting errors and 2. Poisson sampling
𝑝
𝑢𝑛
errors. With regard to the pipetting errors, for a dilution 𝐷 = 𝑞 ≡ (𝑢+𝑣)𝑛 , whereby 𝑢
𝑢
is a volume pipetted in volume 𝑣 to make a dilution of 𝑢+𝑣, and whereby 𝑛 is the step
number in this serially repeated process, the variances of 𝑝, 𝑞, and 𝐷are:
15
20
𝑉𝑎𝑟(𝑝) ≡ 𝑉𝑎𝑟(𝑢𝑛 ) ≅ 𝑛 ∙ 𝑢2(𝑛−1) {𝑉𝑎𝑟(𝑢) + (𝑛 − 1) ∙ 𝐶𝑜𝑣𝑎𝑟(𝑢𝑖 , 𝑢𝑗 )}
(S3)
𝑉𝑎𝑟(𝑞) ≡ 𝑉𝑎𝑟(𝑤 𝑛 ) ≅ 𝑛 ∙ 𝑤 2(𝑛−1) {𝑉𝑎𝑟(𝑤) + (𝑛 − 1) ∙ 𝐶𝑜𝑣𝑎𝑟(𝑤𝑖 , 𝑤𝑗 )}
(S4)
𝑝
𝑉𝑎𝑟(𝐷) ≡ 𝑉𝑎𝑟(𝑞 ) ≅ 𝐷2 {
𝑉𝑎𝑟(𝑝)
𝑝2
+
𝑉𝑎𝑟(𝑞)
𝑞2
−
2𝐶𝑜𝑣𝑎𝑟(𝑝,𝑞)
𝑝𝑞
}
(S5)
whereby 𝑤 = 𝑢 + 𝑣, and indices 𝑖 and 𝑗 are consecutive dilutions. 𝑉𝑎𝑟(𝑢) and
𝑉𝑎𝑟(𝑤) were determined from the (in)accuracy and (im)precision of the pipet, both of
7
which we determined for each Gilson Pipetman used in the preparation of the qPCR
samples. Hedges proofs that the pipetting error, 𝑉𝑎𝑟(𝑃), can be approximated by:
𝑉𝑎𝑟(𝑃) ≅ (𝑢′ )2 ∙ 𝑉𝑎𝑟(𝐷) + 𝐷2 ∙ 𝑉𝑎𝑟(𝑢′ )
(S6)
5
whereby 𝑢′ is the sample volume delivered to the qPCR well. In addition to this serial
dilution variance, due to the randomly distributed nature of DNA marker particles in
each sampling vial, there is a Poisson sampling error, 𝑆, determined as:
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1
𝑉𝑎𝑟(𝑆) = 𝑋 + (𝑢′)2 ∙ 𝑁 ∗ ∙ ∑𝑖=2𝑛
𝑖=𝑛+1{𝑧 𝑖 }
(S7)
whereby 𝑋 is simply the observed DNA marker concentration, 𝑁 ∗ is the estimate of
the unknown original DNA marker concentration, and 1/𝑧 𝑖 is the size of the dilution
step. The first part on the right hand side of eq. (S7) is the error due to the final
15
sampling to be delivered to the qPCR well, and the second part on the right hand side
of eq. (S7) is the error due to the cumulative sampling during the making of the
dilution series. Finally, the confidence interval of a DNA marker concentration at a
95% confidence level due to serial dilution errors can then be determined with:
20
1
1
𝐶. 𝐼. = 1.96 ∙ √𝑉𝑎𝑟(𝑋) ∙ (𝐷)2 ∙ (𝑢′)2
(S8)
whereby:
𝑉𝑎𝑟(𝑋) = 𝑉𝑎𝑟(𝑆) + 𝑉𝑎𝑟(𝑃)
(S9)
8
Random errors
Despite serial dilution errors, identical DNA marker samples should yield identical
threshold cycles, but it is our experience that they do not. In order to quantify the
5
accuracy of the qPCR apparatus, we determined the threshold cycle of 48 identical
samples, and used the standard deviation as a measure for the precision of the
instrument. Finally, in order to quantify the magnitude of the random variation in
threshold cycle from sample taking to qPCR analysis, we prepared a PPCO bottle
with 100 ml of MilliQ water in which we injected 10 µl of concentrated DNA marker
10
solution, and collected duplicate samples every 2 hour for a total duration of 10 hour.
For this case, we used the standard deviation of the DNA marker concentrations as a
measure to quantify the random errors in the entire process from sample taking to
qPCR analysis.
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Results
Assessing confidence intervals of DNA marker concentrations
The (in)accuracy and (im)precision of all pipetting equipment used in our laboratory
was around 2% on a routine basis. With a final sample volume in a PCR well of 4 µl,
the confidence interval from the 1st 10 fold dilution to the 12th 10 fold dilution
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increased from 5 to almost 40% of the initial undiluted concentration (Fig. S1). Since
the standard curve used 10 fold dilutions from step 6 to step 12, meaning that we used
106 to 1012 times serially diluted initial concentration, the confidence interval
associated with the standard curve ranged from 19% to almost 40% of the initial
undiluted concentration. Furthermore, the standard deviation of the qPCR apparatus
25
of 48 identical samples was 0.9% (n=48) of the threshold cycle. Since DNA is
9
multiplied every cycle, a standard deviation of ~1% equalled ~0.1-0.3 cycle, assuming
an amplification protocol of 10-30 cycles. This was equivalent to a DNA mass
standard deviation of 20.1-20.3 or 7-23% of the mean measured concentration. The
standard deviation of 24 samples taken from an identical batch experiment with
5
MilliQ water in order to quantify the random error in the entire process from sample
taking to qPCR analysis, was 14% of the input concentration.
During the tracer injection experiments, we used a 100 times (10·10 fold) serially
diluted sample taken from the PPCO container with concentrated T23 mixed with 100
ml MilliQ water we injected in the brook (see Methods section). The threshold cycle
10
of these samples was around between 10 and 14, while threshold cycles of DNA
marker samples from the brook at measurement point 1 and 2 were between 20 to 26.
A cycle difference of 12 corresponded to a 4-step 10-fold serial dilution of standard
curve samples. So, from DNA input mass at the point of injection to measured DNA
concentration in the brook at the points of measurement involved 6 steps in a 10-fold
15
serial dilution series, which corresponded to a confidence interval of 19.2% of the
input concentration. Together with a random error of 14% of the input concentration,
in our experiments, this gave a confidence interval of ~33% of the input concentration
at a 95% confidence level of a T23 concentration determined in a sample taken from
the field, and such confidence intervals also applied to the batch experiments.
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Amplification efficiencies and standard curve analysis
The Maisbich had a relatively low EC-value (125-135 µS/cm), and low concentrations
of most of the cations (Table S1). The Heuwelerbach had the highest EC-value (412
µS/cm with a relatively high calcium concentration. The Gloop, Bloop, and Ditch had
25
EC-values between 269 and 382 µS/cm, with similar cation concentrations, with,
10
compared to the Luxembourg sites, more Fe and K. Also, the Al concentration in the
Ditch was rather high. Furthermore, in the Gloop and Bloop humic compounds were
present, evidenced by their beer-like appearance. In these latter two streams, none of
the DNA marker material in the positive standard curve samples was amplified,
5
indicating that inhibition of the PCR reaction occurred. For the Gloop, diluting the
samples 4-fold was sufficient to eliminate inhibition, while a 10-fold dilution of the
Bloop samples was sufficient to yield good quality amplification curves. For the other
brooks, no inhibition occurred. We think the inhibition was due to the presence of
humic compounds mentioned above. The standard curves of T22 and T23 (Fig. S2),
10
including the diluted Gloop and Bloop standard curves, were similar for all brooks.
We carried out an analysis of covariances (ANCOVA) of the log-transformed
concentrations versus the threshold cycle, and found that none of the standard curves
were statistically different from each other at a significance level of 5%. The standard
curve efficiency of marker T22 was between 108.4 and 119.4% with an average of
15
113.7%, while for T23, the standard curve efficiency was between 89.4 and 103.8%
with an average of 93.0%. Efficiencies in excess of 100% are somewhat confusing,
and actually indicate that the standard curve is too steep, and that amplification
efficiencies vary for the different standard curve samples. In fact, the individual
amplification efficiencies of T22, determined with LinRegPCR (Fig. S3), ranged
20
between 1.61 and 2.48, while of T23 the individual efficiencies ranged between 1.721.77, and were almost constant over the entire concentration range of the PCR assay.
For marker T22, we also noted random amplification of no template controls at
around 30 cycles of amplification, while for T23 amplification of no template controls
did not occur before a threshold cycle 35 or more. Based on these two assay
25
performance criteria, we concluded that T23 was apparently a better designed DNA
11
marker than T22, although both markers were designed according to the same design
considerations. In order to distinguish positive samples from negative ones, we
applied a difference of 3-5 cycles between the lowest detectable positive sample and
random amplification of a negative control sample. Therefore, for marker T22,
5
injection experiments were considered to be positive when the threshold cycle was
below 25.3-26.9, dependent on the brook. For marker T23, injection experiment
samples were considered to be positive when the threshold cycle was below 31.832.8, dependent on the brook. The T23 concentration belonging to this threshold cycle
was 200 particles in a 20 µl PCR well (Fig. S2), so the lowest possible concentration
10
T23 we could measure in a 4 µl sample was (200/5)·106 = 5·107 particles/L. For
Gloop and Bloop, this minimum measurable concentration had to be multiplied with a
factor of 4 and 10 to account for the dilution that was required to eliminate inhibiting
effects.
15
References
Gaillard, C., and F. Strauss, 2000. Eliminating DNA loss and denaturation during
storage in plastic microtubes. American Biotechnology Laboratory, P. 24, Vol.
18, Issue 13.
Hedges, A.J., 2002. Estimating the precision of serial dilutions and viable bacterial
20
counts. International Journal of Food Microbiology 76 (2002) 207– 214.
Khan, E., and S. Subramania-Pillaib, 2007. Interferences contributed by leaching
from filters on measurements of collective organic constituents. Water
Research, p. 1841-1850, doi: 10.1016/j.watres.2006.12.028.
Ramakers, C., Ruijter, J.M., Lekanne Deprez, R.H., and A. F.M. Moorman, 2003.
25
Assumption-free analysis of quantitative real-time polymerase chain reaction
12
(PCR) data. Neuroscience Letters 339 (2003) 62–66, doi:10.1016/S03043940(02)01423-4
Ruijter, J. M., Ramakers, C., Hoogaars, W. M. H., Karlen, Y., Bakker, O., Van den
Hoff, M. J. B., and A. F. M. Moorman, 2009. Amplification efficiency: linking
5
baseline and bias in the analysis of quantitative PCR data. Nucleic Acids
Research, 2009, Vol. 37, No. 6, Doi:10.1093/nar/gkp045.
13
Tables
Table S1:
Cation concentrations (in mg/L) of the brook waters used in this
research
Stream
5
EC (mS/cm)
Na
Ca
Mg
Al
K
Fe
Maisbich downstream
125
11.5
8.5
4.4
< 0.2
4.0
< 0.2
0.2
-
Maisbich upstream
135
15.7
16.1
5.1
0.3
9.6
0.3
0.4
-
Heuwelerbach
412
2.9
78.7
1.9
0.7
1.1
0.3
0.3
-
Ditch
382
13.2
31.2
7.5
2.8
18.8
1.9
0.2
-
Gelsloopken
345
14.3
31.0
5.2
0.6
17.5
1.3
0.3
yes
Biezenloop
269
12.6
20.3
5.2
0.7
12.1
0.6
0.2
yes
14
Zn Humics
Figures
C.I./ N* (-)
0.4
0.2
0
0
Figure S1:
5
5
10
n-th 10-fold serial dilution (-)
15
Ratio of confidence interval (C.I) at 95% confidence level and undiluted
concentration (N*) as a function of 10-fold serial dilutions for a pipetting
accuracy and (im)precision of 2%, and a final sample volume of 4 µl per
qPCR well
15
Log (part/ml)
8
7
6
5
4
3
2
1
0
milliQ
Heuwelerbach
Maisbich upstream
Ditch
Gloop 4x
Bloop 10x
Log (part/ml)
10
20
25
Threshold cycle
8
7
6
5
4
3
2
1
0
30
35
milliQ
Heuwelerbach
Maisbich upstream
Ditch
Gloop 4x
Bloop 10x
10
Figure S2:
5
15
15
20
25
Threshold cycle
30
35
Standard curves of tracer T22 (above) and T23 (below) for the various
surface waters used in this research. Threshold cycles were determined with
the software of the qPCR device, which is identical to the threshold cycle
algorithm developed by Zhao and Fernald (2005). Data on the y-axis are
expressed as the logarithm of the number of DNA marker particles per µl in a
PCR well, so a value of 1at a threshold cycle of 32.5 in case of T23
10
corresponds to 10 DNA particles per µl or 200 particles in a 20 µl PCR well,
and to (200/4)·106= 5·107 DNA particles per L of sample, since 4 µl of
sample was used per PCR well
16
2.5
T23
T22
Ei
2.0
1.5
1.0
10
Figure S3:
20
Threshold cycle
30
Individual amplification efficiencies, 𝐸𝑖 , determined with LinRegPCR as
function of threshold cycle
5
17