Verification of Principle feasibility
1. Proof of Principle Using ssDNA Substrates
Objective
To verify whether RNase H II can specifically recognize and cleave RNA bases in the probe and the
single base of the target, we ordered single-strand (ssDNA) of 24 nucleotides (nt) in length based
on the MTHFR fragment sequence and used it as the reaction substrate to measure the endpoint
fluorescence intensity.
Procedure
(1) Design of ssDNA substrates: The sequence of the wild-type ssDNA substrate, which is 24 nt long,
corresponds to the MTHFR 677CC, with the 677-site positioned centrally. Similarly, the mutant-type
ssDNA substrate, also 24 nt in length, corresponds to the MTHFR 677TT variant, with the 677-site
located in the middle. The sequence is shown in Table 1.
(2) Design of probes: We designed two allele-specific probes—one targeting the wild-type sequence and the other targeting the mutant MTHFR sequences. The wild-type probe contains an RNA base, rG, which pairs complementarily with the C base in MTHFR 677CC, while the mutant probe contains rA to complement the T base in MTHFR 677TT. Each probe was labeled with both fluorescent and quenching groups: the wild-type probe was labeled with FAM and BHQ1, and the mutant probe was labeled with ROX and BHQ2. The sequence is shown in Table 1.
(3) Construction of the ssDNA-based proof-of-concept reaction system: Prepare 20 µL reaction mixtures for eight groups according to Tables 2 and 3. Incubate the mixtures at 65 °C for 30 min, performing three parallel experiments per group.
(2) Design of probes: We designed two allele-specific probes—one targeting the wild-type sequence and the other targeting the mutant MTHFR sequences. The wild-type probe contains an RNA base, rG, which pairs complementarily with the C base in MTHFR 677CC, while the mutant probe contains rA to complement the T base in MTHFR 677TT. Each probe was labeled with both fluorescent and quenching groups: the wild-type probe was labeled with FAM and BHQ1, and the mutant probe was labeled with ROX and BHQ2. The sequence is shown in Table 1.
(3) Construction of the ssDNA-based proof-of-concept reaction system: Prepare 20 µL reaction mixtures for eight groups according to Tables 2 and 3. Incubate the mixtures at 65 °C for 30 min, performing three parallel experiments per group.
Table 1. The sequences of ssDNA substrates and probes.
The base in red font is the base at MTHFR 677-site.
The /rG/ and /rA/ represent modified RNA bases.
The /rG/ and /rA/ represent modified RNA bases.
Table 2. Experimental system based on ssDNA substrates and wild-type
probe.
The 10×Buffer provided is included with the purchase of RNase H II.
Table 3. Experimental system based on ssDNA substrates and mutant-type
probe.
Results
Figure 1. Proof of principle using ssDNA substrates. A. Fluorescence
intensity based on wild-type probe. B. Fluorescence intensity based on mutant-type probe. Error
bars: SD, n=3
Whether using wild-type or mutant substrates and probes, the fluorescence intensity of the positive
group is significantly higher than that of the control group. This indicates that RNase H II
exhibits excellent specificity in recognition. Preliminary evidence suggests that our experimental
approach, based on the RNase H hypothesis, is feasible.
2. Construction of Plasmids
Objective
Our ultimate goal is to detect genes in blood samples; however, the nucleic acid content in blood is
low, so we need to combine our system with PCR amplification. For this purpose, we designed a
plasmid containing MTHFR gene fragments.
Procedure
(1) We searched for the sequence of MTHFR on NCBI and synthesized corresponding wild-type and mutant
gene fragments with a length of 353 bp at a gene synthesis company.
(2) We separately loaded the DNA fragments into existing plasmid vectors in the laboratory. This plasmid vector is 2694 base pairs (bp) long and confers resistance to ampicillin. The MTHFR gene fragments and vector fragments are shown in the Figure 2.
(3) After introducing the recombinant plasmids into DH5α competent cells, we cultured the cells and selected single colonies for expansion. The image of the agar plate containing bacterial colonies is shown in the Figure 3.
(3) We then extracted plasmids from the enriched bacterial cultures and sequenced them. Upon successful sequencing, these two plasmids were used as templates for subsequent experiments.
(2) We separately loaded the DNA fragments into existing plasmid vectors in the laboratory. This plasmid vector is 2694 base pairs (bp) long and confers resistance to ampicillin. The MTHFR gene fragments and vector fragments are shown in the Figure 2.
(3) After introducing the recombinant plasmids into DH5α competent cells, we cultured the cells and selected single colonies for expansion. The image of the agar plate containing bacterial colonies is shown in the Figure 3.
(3) We then extracted plasmids from the enriched bacterial cultures and sequenced them. Upon successful sequencing, these two plasmids were used as templates for subsequent experiments.
Results
Figure 2. 1% agarose gel diagram. Lanes 1 and 3 contain amplified plasmid
vectors, while Lanes 2 and 4 contain the MTHFR target fragment. The former corresponds to the
wild type, and the latter corresponds to the mutant type.
Connect the MTHFR fragment to the carrier skeleton, transfer the reaction product into DH5α
competent cells, and culture them on solid medium. The results are shown in Figure 3.
Figure 3. Solid culture medium plate with bacterial colonies. On the left
are the colonies grown after introducing the wild-type plasmid into DH5α competent cells, and on
the right are the colonies grown after introducing the mutant plasmid into DH5α competent cells.
Single colonies were picked from two solid plates for enrichment culture in liquid medium, after
which plasmids were extracted for agarose gel electrophoresis. The results are shown in Figure 4.
The plasmid were also sequenced, and partial sequencing results are shown in Figure 5.
Figure 4. 1% agarose gel diagram. Lane1: MTHFR target fragment; Lane 2:
Vector; Lane 3: wild-type plasmid; Lane 4: mutant-type plasmid.
Figure 5. Partial screenshot of sequencing results. The red letters on
the graph correspond to the base of MTHFR 677-site.
According to Figure 4 and 5, it can be concluded that the MTHFR fragment has been successfully
inserted into the plasmid vector, resulting in a complete plasmid. The wild-type and mutant plasmids
will be used as templates for subsequent experimental studies.
3. Proof of Principle Using Plasmids
The experimental validation of the ssDNA substrates has been successfully completed. To determine
whether this system is compatible with the PCR amplification system, we conducted subsequent
experiments using plasmids as reaction templates. We designed reaction systems incorporating both
wild-type and mutant plasmids, along with associated reaction components. We monitored real-time
fluorescence curves to verify the feasibility of the experimental principle based on the
fluorescence data.
3.1 Verification of Agarose Gel Electrophoresis
Procedure
Procedure
(1) Design of sequence: The probe sequence is the same as Table 1. The PCR primer sequences are
shown in Table 4. The length of the amplification product generated by this pair of primers is 191
bp.
Table 4. The sequences of primers.
(2) Perform PCR amplification reaction according to the Table 5.
(3) The reaction products were analyzed using agarose gel electrophoresis, and amplification was assessed based on the presence and intensity of the bands.
(3) The reaction products were analyzed using agarose gel electrophoresis, and amplification was assessed based on the presence and intensity of the bands.
Table 5. The reaction system of agarose gel electrophoresis experiment.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40 cycles
Results
Figure 6. 3% Agarose gel image for plasmid verification.
The length of the amplification product generated by this pair of primers is 191 bp, which is
consistent with the position shown in the gel image. This indicates successful amplification, and
the designed primers can be used for subsequent experiments.
3.2 Verification of Real-Time Fluorescence Curves
We proceeded with qPCR experiments using a single probe in one-tube. The reaction system is shown in
Table 6. The success of the experiment was determined based on the output fluorescence signal.
Table 6. The reaction system of qPCR.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Results
Figure 7. Real-time fluorescence curves. A. Fluorescence curve of the
Wild-type group. B. Fluorescence curve of the Mutant-type group.
When plasmids, probes of the same type as the plasmids, and RNase H II are present simultaneously in
the system, the real-time fluorescence curve shows a significant upward trend. In the absence of
RNase H II enzyme, the curve remains flat, and the △Rn value is zero, indicating that RNase H is
essential and compatible with the PCR system. The fluorescence signal accurately reflects the
amplification of plasmids. Our experimental feasibility has been confirmed.
Reflection
The combined results from experiments of ssDNA substrates and plasmids largely align with
expectations, demonstrating that wild-type and mutant type genes of MTHFR can be detected under
laboratory conditions.
Optimization of Condition
1. Optimization of operational steps
Objective
The single-probe in single-tube detection method cannot accurately distinguish the CC, CT, and TT
genotypes in a single experiment. To determine a sample's genotype, two separate tubes must be
tested, each containing a different probe. The genotype is then inferred by comparing the distinct
fluorescence signals (FAM and ROX) emitted by each tube. This process is cumbersome and inefficient.
To achieve the technology of dual-probe in one-tube detection and improve both efficiency and operational convenience, subsequent experiment aims to verify the feasibility of using wild-type and mutant probes simultaneously within a one tube.
To achieve the technology of dual-probe in one-tube detection and improve both efficiency and operational convenience, subsequent experiment aims to verify the feasibility of using wild-type and mutant probes simultaneously within a one tube.
Procedure
Table 7. The qPCR reaction system with dual-probes in one-tube.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Results
Figure 8. Dual-probe in one-tube detection. The positive groups in the
chart of A, B and C correspond to group 1, 2 and 3 in Table 1, while the control group
corresponds to group 4, 5 and 6. A. Using wild plasmids as amplification templates. B.
Amplification templates are mutant plasmids. C. Amplification templates are both wild-type and
mutant plasmids. The shaded area in the figures indicates the error bar. Error bars: SD, n=3.
When two probes were added to each tube, the following observations were made: if the template
contained only wild-type plasmids, only the FAM fluorescence signal from the wild-type probes showed
a significant increase; if the template contained only mutant plasmids, only the ROX fluorescence
signal from the mutant probes showed a significant increase; and if both wild-type and mutant
plasmids were present, significant increases were observed in both the FAM and ROX signals. The
control groups without RNase H II in each case showed no increase in ΔRn, indicating that RNase HII
is essential for this experiment.
Consideration
The experimental results indicate that the dual-probes, one-tube method can effectively distinguish
between wild-type and mutant variants, offering greater simplicity in detection procedures. After
this experiment, subsequent studies adopted a dual-probe, one-tube detection strategy.
In order to facilitate the selection of optimal experimental conditions, we simulated heterozygous blood by using both wild-type and mutant plasmids as reaction templates to test their fluorescence responses to two probes in subsequent experiments. When the real-time curves of two fluorescence signals tend to coincide, it is the optimal condition we need to select. For this purpose, we have introduced the evaluation criteria of ΔΔRn. The ΔΔRn value is equal to the absolute value of the ΔRn of FAM signal and the ΔRn of ROX signal. The smaller the value of ΔΔRn, the higher the overlap between the two curves.
In order to facilitate the selection of optimal experimental conditions, we simulated heterozygous blood by using both wild-type and mutant plasmids as reaction templates to test their fluorescence responses to two probes in subsequent experiments. When the real-time curves of two fluorescence signals tend to coincide, it is the optimal condition we need to select. For this purpose, we have introduced the evaluation criteria of ΔΔRn. The ΔΔRn value is equal to the absolute value of the ΔRn of FAM signal and the ΔRn of ROX signal. The smaller the value of ΔΔRn, the higher the overlap between the two curves.
2. Optimization of Primer Sequences
Objective
We designed three different sets of primers and selected the optimal primer through experimental
testing. Primer sequences are shown in Table 8.
Table 8. The Primer Sequences.
Procedure
Table 9. The qPCR systems in three primer sequences.
*The primers used in groups 1, 2, and 3 correspond to the first, second,
and third sets of primers listed in Table 9, respectively.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40 cycles
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40 cycles
Results
Figure 9. Optimization of primer sequences. A. Real time fluorescence
curve of the primer-1. B. Real time fluorescence curve of the primer-2. C. Real time
fluorescence curve of the primer-3. D. ΔRn value of the 40th cycle. The ΔΔRn value is equal to
the absolute value of the ΔRn of FAM signal and the ΔRn of ROX signal. Error bars: SD, n=3.
The first set of primers performed well, so it was chosen as the primer for subsequent experiments.
3. Optimization of RNase H II Dosage
Objective
To determine the optimal RNase H II dosage in qPCR reactions for improved efficiency and accuracy in
detecting wild-type and mutant plasmids, we designed reaction systems with varying RNase H II
concentrations. Experiments strictly controlled for single variables, maintaining consistent
reaction components and conditions while varying only RNase H II dosage. Three gradient
concentrations (0.1 µL, 0.25 µL, 0.5 µL) were established, with corresponding reaction setups for
each gradient.
Procedure
(1) Place prepared qPCR reaction mixtures into the qPCR instrument for reaction.
(2) Record the fluorescence curves for each reaction group.
(3) Process the data to obtain FAM and ROX values at different RNase H II dosage.
(2) Record the fluorescence curves for each reaction group.
(3) Process the data to obtain FAM and ROX values at different RNase H II dosage.
Table 10. The qPCR reaction system for optimizing of RNase H II dosage.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Results
Figure 10. Optimization of RNase H II Dosage. A. The dosage of RNase H II
is 0.1 µL. B. The dosage of RNase H II is 0.25 µL. C. The dosage of RNase H II is 0.5 µL. D. ΔRn
value of the 40th cycle. The ΔΔRn value is equal to the absolute value of the ΔRn of FAM signal
and the ΔRn of ROX signal. Error bars: SD, n=3.
The strong fluorescence signal was observed when 0.25 µL of RNase H II was added, and the ΔΔRn value
is the smallest, indicating that 0.25 µL is the optimal amount.
4. Optimization of Probe Concentration and Proportion
Objective
To determine the optimal concentration ratio of wild-type and mutant probes in qPCR reactions,
thereby enhancing fluorescence detection efficiency and accuracy. Reaction systems with varying
probe concentrations were designed. This experiment compared four probe concentration ratios.
The experiment strictly controlled for single variables, maintaining consistent reaction components and conditions while varying only the Wild-type and Mutant probe volumes. Three gradient ratios were set: 0.5 µL:1 µL,1 µL:1 µL, 2 µL:1 µL, and 4 µL:1 µL. Each ratio underwent corresponding reaction setups to compare the impact of different probe volumes on qPCR outcomes.
The experiment strictly controlled for single variables, maintaining consistent reaction components and conditions while varying only the Wild-type and Mutant probe volumes. Three gradient ratios were set: 0.5 µL:1 µL,1 µL:1 µL, 2 µL:1 µL, and 4 µL:1 µL. Each ratio underwent corresponding reaction setups to compare the impact of different probe volumes on qPCR outcomes.
Procedure
Table 11. The qPCR reaction system for optimizing of probe concentration.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Results
Figure 11. Optimization of probe concentration and proportion. A. 0.5 μL of
10 μM wild-type probe and 1 μL of 10 μM mutant probe were added. B. 1 μL of 10 μM wild-type probe
and 1 μL of 10 μM mutant probe were added. C. 2 μL of 10 μM wild-type probe and 1 μL of 10 μM mutant
probe were added. D. 4 μL of 10 μM wild-type probe and 1 μL of 10 μM mutant probe were added. E. ΔRn
value of the 40th cycle. The ΔΔRn value is equal to the absolute value of the ΔRn of FAM signal and
the ΔRn of ROX signal. Error bars: SD, n=3.
Adding 2 μL of 10 μM wild-type probe and 1 μL of 10 μM mutant probe constituted the optimal condition.
Analytical Performance
1. Specificity
To test the specificity of the system, we selected plasmids containing BRAF gene fragments, whose
vectors were the same as that of the MTHFR plasmid. The BRAF gene is a proto-oncogene located on the
long arm of human chromosome 7 (7q34). V600E mutation is the most common SNP mutation in the BRAF
gene,
which is associated with thyroid cancer and melanoma. It results from a substitution of the thymine
(T)
base with adenine (A) at position 1799, leading to the replacement of valine with glutamic acid at
amino
acid position 600. Therefore, we used wild-type and mutant BRAF plasmids as reaction templates,
keeping
the other components of the system unchanged, and judged the specificity of our scheme based on the
response of the fluorescence signal.
Table 12. The qPCR reaction system for specificity experiment.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Results
Figure 12. The specificity experiment. A. Real time fluorescence curve.
B.
ΔRn value of the 40th cycle. Error bars: SD, n=3.
When the template is BRAF plasmid, there is no upward trend in the fluorescence signals of FAM and
ROX.
This result demonstrates that our detection method has excellent specificity for the MTHFR gene.
2. Sensitivity
Objective
To determine the sensitivity of the detection system, specifically calculating its detection limit
and
precision, we designed reaction systems with varying plasmid concentrations. This experiment
compared
six plasmid concentrations. The experiment strictly controlled for single variables, maintaining
consistent reaction components and conditions while varying only the wild-type and mutant plasmid
quantities. Six concentration gradients (10⁶, 10⁵, 10⁴, 10³, 10², 10 copies/µL) were established to
investigate the plasmid concentrations distinguishable by the detection system.
Ct (Cycle threshold) is the cycle number at which the fluorescence signal in real - time PCR first crosses a preset threshold. It’s inversely related to the initial amount of target nucleic acid; lower Ct means higher initial concentration. Therefore, Ct values are also used as our presentation results.
Ct (Cycle threshold) is the cycle number at which the fluorescence signal in real - time PCR first crosses a preset threshold. It’s inversely related to the initial amount of target nucleic acid; lower Ct means higher initial concentration. Therefore, Ct values are also used as our presentation results.
Procedure
Table 13. The qPCR reaction system for sensitivity experiment with wild
plasmid.
Table 14. The qPCR reaction system for sensitivity experiment with wild
plasmid.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Results
Figure 13. The sensitivity of the assay. A. Real-time fluorescence curve
of
wild-type plasmid. B. The Ct values of different concentrations of wild plasmids. C. Real-time
fluorescence curve of mutant-type plasmid. D. The Ct values of different concentrations of
mutant-type plasmids. Error bars: SD, n=3.
According to the real-time fluorescence curve results (Figure 13A and Figure 13C), when the plasmid
concentration is below 10^3 copies/μL, the curve cannot be distinguished from that of lower
concentrations. Therefore, we determine the sensitivity to be 10^4 copies/μL. As the plasmid
concentration decreases, the Ct value increases (Figure 13B and Figure 13D), which meets the
expected
results.
Blood Samples Experiments
1. Agarose Gel Electrophoresis for Amplification Proof
Objective
To verify that the genes in the blood sample direct amplification are identical to those in the
plasmid
experiment.
Procedure
(1) Amplification using blood samples as templates.
(2) Gel Electrophoresis.
(2) Gel Electrophoresis.
Results
Figure 14. Agarose Gel Electrophoresis. Gel images of blood and plasmid
amplification.
As shown by the gel image, the blood group exhibits the same gene length as the plasmid group.
Serving
as a control for the plasmid group, the blood group confirms the successful insertion of the MTHFR
gene
into the plasmid.
2. Principle verification based on blood samples
2.1 Two-step amplification method
Objective
Considering the low nucleic acid content in blood, PCR amplification is initially performed on the
blood
sample, and the amplified product is subsequently used as the template for qPCR.
Procedure
Table 15. The PCR reaction system.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Table 16. The qPCR reaction system.
Reaction procedure: 94°C, 5min + ( 94°C,30s + 55°C, 30s + 68°C,25s)×40
cycles
Result
Figure 15. Real time fluorescence curve of qPCR step. A. Real-time
fluorescence curve of wild-type sample in the step of qPCR. B. Real-time fluorescence curve of
heterozygote sample in the step of qPCR. Error bars (Shaded area): SD, n = 3.
As shown in the Figure 15, the FAM fluorescence signal curve of the wild-type sample showed the most
significant increase, while the FAM and ROX fluorescence curves of the heterozygous sample both
showed a
significant increase, indicating that the feasibility of the two-step method was passed. The
vertical
axis of the experimental results is represented by Rn instead of ΔRn because the fluorescence
increases
too rapidly, making it unsuitable for baseline subtraction.
The experimental results of the two-step method demonstrated the feasibility of detecting blood samples. Due to the rapid amplification speed of the two-step method and the lack of a baseline, we attempted to use the one-step method for blood samples.
The experimental results of the two-step method demonstrated the feasibility of detecting blood samples. Due to the rapid amplification speed of the two-step method and the lack of a baseline, we attempted to use the one-step method for blood samples.
2.2 One-step amplification method
Objective
To validate the feasibility of direct blood amplification experiments, specifically to determine
whether
the real-time fluorescence curve of qPCR from direct blood amplification can accurately identify
gene
variants related to folate metabolism, we conducted a one-tube reaction.
Procedure
Add 100 µL of blood sample to 500 µL of red blood cell lysis buffer. Mix by inverting the tube and
incubate at 4°C for 20 minutes. After incubation, centrifuge at 5000 rpm for 5 minutes and discard
the
supernatant. Resuspend the pellet in 50 µL of red blood cell lysis buffer. Use 5 µL of the
resuspension
as the reaction template.
Table 17. The qPCR reaction system for blood in one-step.
94℃,5min + ( 94℃,30s + 55℃,30s + 68℃,25s) ×45 cycles
Results
Figure 16. Real-time fluorescent curves of qPCR in one-tube. A. Real-time
fluorescence curve of heterozygous blood samples. B. Real-time fluorescence curve of wild-type
blood
samples. C. Real-time fluorescence curve of mutant blood samples. Error bars (Shaded area): SD,
n =
3.
The results of the blood samples are the same as those of the plasmid samples: if the template
contained
only wild-type blood, only the FAM fluorescence signal from the wild-type probes showed a
significant
increase; if the template contained only mutant blood, only the ROX fluorescence signal from the
mutant
probes showed a significant increase; and if the template contained only heterozygous blood,
significant
increases were observed in both the FAM and ROX signals.
qPCR results for wild-type, heterozygous, and mutant variants all matched sequencing data, confirming that the system can be successfully applied to the detection of actual blood samples.
qPCR results for wild-type, heterozygous, and mutant variants all matched sequencing data, confirming that the system can be successfully applied to the detection of actual blood samples.
3. Optimization based on blood samples
3.1 Optimization of Blood Incubation Time
Objective
Objective
Determine the optimal incubation time. Establish an accurate, rapid, and efficient detection system
that
yields stronger fluorescence signals while ensuring the accuracy and reproducibility of results.
Procedure
Using heterozygous blood as the reaction template, establish three distinct incubation time
gradients
for the reaction: 10 min, 20 min, 30 min.
Results
Figure 17. Blood Incubation Time Optimization. A. Real-time fluorescence
curve of incubation condition of 10 min. B. Real-time fluorescence curve of incubation condition
of
20 min. C. Real-time fluorescence curve of incubation condition of 30 min. D. ΔRn value of the
40th
cycle. The ΔΔRn value is equal to the absolute value of the ΔRn of FAM signal and the ΔRn of ROX
signal. Error bars: SD, n = 3.
The ΔRn values for FAM and ROX in the 10-minute fluorescence curve were around 50000. Extending the
incubation time to 20 minutes increased ΔRn to approximately 80000. However, further extending the
incubation time to 30 minutes did not result in a significant change in ΔRn. Though the value of
ΔΔRn in
10 min-incubation condition was the smallest, its ΔRn is weaker than under 20 min incubation
conditions.
Therefore, 20 min was the most favorable incubation condition and it was selected as the optimal
incubation time for the red blood cell lysis buffer.
3.2 Optimization of Template Quantity
Objective
Objective
Determine the optimal template quantity. To complete the assay using minimal blood samples while
ensuring accurate and reproducible results.
Procedure
Maintaining a constant total qPCR volume, we designed blood sample loading volumes of 2 μL, 5 μL,
and 7
μL. The system was then top-up to 25 μL with ddH₂O. To ensure experimental accuracy and
reproducibility,
we selected heterozygous blood sample and performed three replicate experiments.
Results
Figure 18. Template Quantity Optimization. A. Real-time fluorescence
amplification curve of 2 μL template quantity. B. Real-time fluorescence amplification curve of
5 μL
template quantity. C. Real-time fluorescence amplification curve of 7 μL template quantity. D.
Comparison of different template volume bar charts. The ΔRn value corresponds to the 40th cycle.
Error bars: SD, n = 3.
Both Wild-type and Mutant probes exhibited strong fluorescence at 5 µL template volume. Compared to
the
2 µL and 7 µL groups, the ΔRn values of 40th cycle was stronger at 5 µL. Thus, 5 µL was selected as
the
optimal template volume.
Reflection
Analysis of the phenomenon where the 7 µL group exhibited lower fluorescence intensity than the 5 µL
group revealed that the composition of the red blood cell lysate affected polymerase activity. The 7
µL
group contained a higher volume of red blood cell lysate, and the salt solution (EDTA) within the
lysate
inhibited enzyme activity, resulting in reduced fluorescence intensity in the 7 µL group.
Application of POCT
RPA Overview
RPA technology is a novel isothermal nucleic acid amplification technique. Its core principle
involves
the formation of a dynamic complex between a recombinase and specific primers. This complex actively
scans double-stranded DNA, mediating target sequence recognition and strand invasion. With the
synergistic action of single-stranded DNA-binding proteins, the complex can stably unwind
single-stranded templates while preventing their re-annealing, thereby replacing the
high-temperature
denaturation step in traditional PCR. Subsequently, DNA polymerase with strand displacement activity
continuously extends primers at a constant temperature of 37-42°C, achieving exponential nucleic
acid
amplification. This mechanism enables RPA reactions to complete amplification within 15-20 minutes
in a
constant-temperature environment without requiring sophisticated equipment like thermal cyclers. It
offers advantages of speed, sensitivity, and portability, aligning with the project's objective of
developing rapid test strips.
RNase H II Overview:
For POCT applications, we propose distinguishing CT, TT, and CC genotype patients by designing
Mutant
and Wild-type probes and measuring fluorescence intensity during RPA amplification. Probes were
engineered for wild-type and mutant MTHFR variants to bind their respective targets. The Wild-type
probe
features a FAM fluorescent group at 5’ end and a quencher group BHQ1 at 3’ end, while the Mutant
probe
carries a ROX fluorescent group at 5’ end and a quencher group BHQ2 at 3’ end. RNase H II
specifically
recognizes DNA-RNA hybrid double strands at its optimal temperature. Upon binding to the probe, it
cleaves the probe, separating the fluorescent group from the quencher group, thereby emitting
fluorescence. This allows us to determine the patient's genotype by measuring fluorescence values.
1. Amplification Proof
Objective
To achieve one-tube amplification of the target MTHFR gene in vitro, we employ RPA technology.
Correct
primer-target gene pairing is critical for successful amplification. We designed four primer pairs
(Table. 18) and conducted experiments to select the most specific primers with the clearest
amplification results, laying the foundation for subsequent experiments.
Procedure
Table 18. The different primers for RPA.
Results
Figure 19. Gel electrophoresis patterns of amplification products with
different
Figure 20. Gel electrophoresis patterns of amplification products with
primer
pair 3.
The gel electrophoresis image of the RPA products with different primers are shown in the Figure 19.
It
is evident that primer pair 3 exhibits the most pronounced amplification result. We also observe
very
faint bands appearing in the blank control group for each primer set. To rule out the influence of
aerosols in the air, we conducted a separate validation specifically for primer pair 3 (Figure 20).
Ultimately, it was confirmed that primer pair 3 is the most suitable primer pair for the RPA
experiment.
To further verify the success of the amplification, we added SYBR Green I dye to the RPA system.
SYBR
Green I binds to double-stranded DNA and emits fluorescence, allowing the amplification status to be
monitored through a real-time fluorescence curve.
Figure 21. Real-time fluorescence curve with SYBR Green I.
According to Figure 21, the fluorescence of the positive group containing plasmids showed a
significant
increase, while the control group without plasmids did not show a significant increase in
fluorescence.
Consideration
During the experiment, we observed that the RPA system exhibits high sensitivity, primarily
manifested
in the inevitable appearance of non-specific bands in the blank group regardless of conditions. This
places stringent demands on our experimental environment. Additionally, we can mitigate
environmental
stress by designing primers with enhanced specificity. Although airborne aerosols may influence the
reaction, RPA amplification was successful. This indicates that during exponential amplification,
the
impact of these aerosols is negligible.
2. Feasibility verification of RPA System
Objective
RPA reactions typically occur at 38°C, while RNase H II cleavage activity temperature is between
50-70
°C. To simplify the experimental setup and facilitate future test strip design, we need to identify
an
optimal temperature where the RPA-RNase H II system generates distinct fluorescent signals, enabling
differentiation between blood samples of different genotypes.
Procedure
We set up different temperature gradients for PCR experiments, and the fluorescence intensity in
each
reaction system was collected every minute during the experimental process. Incubate at 38 °C for 30
min
initially, then adjust the temperature to 50 °C, 60 °C, and 70 °C, incubating for 30 min at each
temperature.
Three parallel sets of each experiment were performed. In order to better present the fluorescence trend, we have omitted the error bars.
Three parallel sets of each experiment were performed. In order to better present the fluorescence trend, we have omitted the error bars.
Result
Figure 22. Real-time fluorescence amplification curves for 30 min of
amplification at 38 °C followed by 30 min at 50 °C (cycles 1 to 30 at 38 °C and cycles 31 to 60
at
50 °C, with each cycle lasting 1 minute). Both FAM and ROX fluorescence channels were monitored
simultaneously.
Figure 23. Real-time fluorescence amplification curves for 30 min of
amplification at 38 °C followed by 30 min at 60 °C (cycles 1 to 30 at 38 °C and cycles 31 to 60
at
50 °C, with each cycle lasting 1 minute). Both FAM and ROX fluorescence channels were monitored
simultaneously.
Figure 24. Real-time fluorescence amplification curves for 30 min of
amplification at 38 °C followed by 30 min at 70 °C (cycles 1 to 30 at 38 °C and cycles 31 to 60
at
50 °C, with each cycle lasting 1 minute). Both FAM and ROX fluorescence channels were monitored
simultaneously.
The results of all three groups of experiments did not meet our expectations. The fluorescence
intensity
of each group did not show a significant increase during the amplification process; on the contrary,
the
initial value of fluorescence intensity obtained was high and remained stable before the temperature
change, and the fluorescence intensity decreased after the temperature increase.
Thoughts
We hypothesize that the probe binds only to single-stranded DNA, whereas isothermal amplification
yields
double-stranded DNA. Temperatures of 50/60°C are insufficient to unwind the double strands and allow
probe binding. Additionally, single-stranded DNA binding proteins within the RPA system may also
interfere with probe binding.
Objective
To verify whether double-stranded DNA impedes probe binding, we attempted high-temperature
pretreatment
after RPA to denature DNA, single-strand binding proteins, or other RPA system components that may
hinder probe binding.
Procedure
Program: 38°C, 30 min → 90°C, 10 min → 70°C, 30 min
Results
Figure 25. Multicomponent Plot
Figure 26. ΔF histogram. ΔF =Fluorescence intensity at 70th cycle -
fluorescence intensity at 40th cycle. Error bars: SD, n = 3.
It is speculated that after incubation at 95°C, the substance affecting RNase H II in the RPA system
was
inactivated. However, the system underwent two temperature changes, increasing its complexity.
Furthermore, the high temperatures required for the probe reaction (70°C) and the inactivation of
the
RPA system (90°C) are difficult to achieve under test strip conditions. We should reconsider the
selection and optimization of detection and identification techniques.
3. The effect of RNase H II enzyme dosage
Objective
In preliminary experiments, we observed that the initial fluorescence readings in the RPA assay were
unusually high, and the fluorescence curve trends between the experimental and blank groups showed
little difference—contrary to our expectations. We suspect that an excessively high RNase H II
enzyme
concentration caused the enzyme to cleave most probes during the initial amplification phase of RPA,
resulting in premature fluorescence emission. Therefore, we decided to reduce the RNase H II enzyme
concentration and explore other potential factors.
Procedure
Dilute RNase H II enzyme concentration tenfold.
Results
Figure 27. Amplification curve at 38°C (38°C for 1 to 30 cycles, each
cycle
lasting 1 min). The top red line represents the ROX positive control group.
Figure 28. Real-time fluorescence amplification curve of RPA (38°C for 1
to
30 cycles and 60°C for 31 to 60 cycles, each cycle lasting 1 min).
We observed that the fluorescence intensity was higher in the system without RNase H II than in the
system with RNase H II. Furthermore, the fluorescence growth trend was nearly identical regardless
of
RNase H II presence. The experimental feasibility of RPA and RNase H II has still not been approved.
Reflection
The experimental results deviated significantly from our expectations. This also indicates that the
increase in fluorescence intensity is largely independent of RNase H II activity. We suspect this
may be
due to the inherently high background fluorescence of the probe itself, coupled with interference
from
other enzymes in the selected RPA kit, which may hinder probe binding to the target gene fragment
and
partially inactivate RNase H II. In addition, we believe that the significant decrease in
fluorescence
intensity after warming may be due to the annealing of the probe during warming, resulting in the
fluorescent group being close to the bursting group and the fluorescence intensity being weakened.
We
will conduct further experiments with variable adjustments to investigate the specific cause.
4. Further explore the feasibility of the experiment
Objective
Based on prior experiments, we observed that the fluorescence intensity curves for both wild-type
and
mutant variants obtained during optimization did not align with expectations. The fluorescence curve
trends between experimental and blank groups showed minimal difference, precluding subsequent
differentiation. Therefore, we designed the next experiment to run the RPA system and our RNase H II
system separately, verifying whether the RPA system successfully amplified.
Results
(1) RPA system results:
Figure 29. Gel images of RPA amplification results
(2) Fluorescence curve obtained after introducing the RNase H II system into the reaction：
Figure 30. Real-time fluorescence amplification curve of RPA (50 °C for 1
to
60 cycles, each cycle lasting 1 min).
Experimental results indicate that amplification in the RPA system at 38°C was successful. However,
subsequent introduction of the RNase H II system still yielded no significant difference in
fluorescence
curve trends between the experimental group and the blank group.
Reflection
After consecutive setbacks in temperature optimization and RNase H II enzyme concentration
optimization,
we hypothesized that components other than RNase H II within the reaction system were degrading the
probe. This likely caused the failure of the post-RPA fluorescence intensity measurement protocol.
With this question in mind, we consulted the kit manufacturer and reviewed relevant literature. Ultimately, we discovered that DNA polymerase I used in the RPA kit possesses 5'-3' exonuclease activity. During the amplification phase, single-stranded binding proteins in the RPA system attach the probe to the front of the primer. However, DNA polymerase I cleaves the probe bound to the primer during amplification, prematurely releasing fluorescence. This incompatibility between RPA and RNase H II systems necessitates modifying the entire system for point-of-care testing (POCT) applications.
With this question in mind, we consulted the kit manufacturer and reviewed relevant literature. Ultimately, we discovered that DNA polymerase I used in the RPA kit possesses 5'-3' exonuclease activity. During the amplification phase, single-stranded binding proteins in the RPA system attach the probe to the front of the primer. However, DNA polymerase I cleaves the probe bound to the primer during amplification, prematurely releasing fluorescence. This incompatibility between RPA and RNase H II systems necessitates modifying the entire system for point-of-care testing (POCT) applications.
