Our goal is to detect single-base mutations in human blood genes. After discussions with the principal investigator and instructor, we developed a preliminary project design plan. First, we will verify the principle using a single-stranded substrate containing the target site. Next, we will construct a plasmid containing the target gene fragment to serve as a reaction template for validating the experiment's feasibility. After confirming feasibility, we will conduct performance optimization experiments. Finally, the method will be applied to actual human blood samples for testing.
We have conducted the DBTL cycle in all aspects of the project and will explain how to apply these loops to our experimental design.
Principle Verification Based on ssDNA Substrate
Design
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.
(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.
(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.
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.
Figure 1. Schematic Diagram of an Experiment Based on the ssDNA Principle.
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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 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.
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Figure 2. 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
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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.
Principle Verification Based on Plasmid
Cycle 1: Construction of plasmids
Design
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.
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(1) We searched for the sequence of MTHFR on NCBI and selected a fragment containing the target gene. Then we synthesized corresponding wild-type and mutant gene fragments at a gene synthesis company, with a length of 353 bp.
(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.
(3) After introducing the recombinant plasmids into DH5α competent cells, we cultured the cells and selected single colonies for expansion.
(4) 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.
(3) After introducing the recombinant plasmids into DH5α competent cells, we cultured the cells and selected single colonies for expansion.
(4) 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.
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Figure 3. The plasmid profile of MTHFR mutant-type plasmid.
Figure 4. The plasmid profile of MTHFR wild-type plasmid.
Figure 5. 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.
Figure 6. Partial screenshot of sequencing results in wild-type plasmid. The red letters on the graph correspond to the base of MTHFR 677-C.
Figure 7. Partial screenshot of sequencing results in mutant-type plasmid. The red letters on the graph correspond to the base of MTHFR 677-T.
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The sequencing results are consistent with our expectations. Therefore, the plasmid loaded with MTHFR gene fragments was used as reaction templates to participate in subsequent experiments.
Cycle 2: Principle Verification with single-probe in single-tube
Design
To verify the specificity of fluorescent probes in recognizing wild-type and mutant plasmids in qPCR reactions and to investigate the catalytic role of RNase H II, we designed reaction systems containing different plasmid combinations and RNase H II. When a probe binds to its corresponding plasmid template, RNase H II cleaves the probe, separating the fluorophore from the quencher and generating fluorescence. By monitoring the real-time fluorescence curves, the specificity of the probes and the enzymatic activity can be evaluated.
The experiment consisted of six reaction groups, with variables being the presence or absence of wild-type plasmid, mutant plasmid, and RNase H II, while keeping other reaction components consistent. This design aimed to clarify the effects of different plasmid combinations and RNase H II on fluorescence signals.
The experiment consisted of six reaction groups, with variables being the presence or absence of wild-type plasmid, mutant plasmid, and RNase H II, while keeping other reaction components consistent. This design aimed to clarify the effects of different plasmid combinations and RNase H II on fluorescence signals.
Figure 8. Schematic diagram of the single probe in single-tube reaction. The fluorescence signal of the wild-type probe is FAM, while the fluorescence signal of the mutant probe is ROX. Both signals can be monitored simultaneously.
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(1) Design of sequence: 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.
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
Then, 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
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Figure 9. 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.
Figure 10. 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 ΔRn 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.
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The results demonstrate that SNP of MTHFR genes can be detected under laboratory conditions. The principle has been preliminarily validated successfully.
Optimization of Condition
Cycle 1: Optimization of operational steps
Design
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 inferred by comparing the distinct fluorescence signals (FAM and ROX) emitted by each tube. This process is cumbersome and inefficient.
Therefore, achieving the technology of dual-probe in one-tube detection is essential. This experiment aims to verify the feasibility of using wild-type and mutant probes simultaneously within a one tube.
Therefore, achieving the technology of dual-probe in one-tube detection is essential. This experiment aims to verify the feasibility of using wild-type and mutant probes simultaneously within a one tube.
Figure 11. Schematic diagram of the dual probe in single-tube reaction. The fluorescence signal of the wild-type probe is FAM, while the fluorescence signal of the mutant probe is ROX. Both signals can be monitored simultaneously.
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Prepare the reaction solution according to the system outlined in Table below, and place the solution into an 8-tube strip and put it into a qPCR machine for reaction. Simultaneously monitor the fluorescence signals of both FAM and ROX dyes for each tube.
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
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Figure 12. 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 figure 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.
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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.
Cycle 2: Optimization of Primer Sequences
Design
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.
We designed three different sets of primers and selected the optimal primer through experimental testing. Primer sequences are shown in Table 8.
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.
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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 X, 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
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Figure 13. 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.
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The first set of primers performed well, so it was chosen as the primer for subsequent experiments.
Cycle 3: Optimization of RNase H II Dosage
Design
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.
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(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
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Figure 14. 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.
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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.
Cycle 4: Optimization of Probe Concentration and Proportion
Design
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.
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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.
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
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Figure 15. 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.
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Adding 2 μL of 10 μM wild-type probe and 1 μL of 10 μM mutant probe constituted the optimal condition.
Analytical Performance
Cycle 1: Testing the specificity of the system
Design
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.
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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
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Figure 16. The specificity experiment. a, Real time fluorescence curve. b, ΔRn value of the 40th cycle. Error bars: SD, n=3.
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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.
Cycle 2: Testing the sensitivity of the system
Design
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, 101 copies/µL) were established to investigate the plasmid concentrations distinguishable by the detection system.
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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.
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
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Figure 17. 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.
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According to the real-time fluorescence curve results (Figure 2A and 2C), when the plasmid concentration is below 103 copies/μL, the curve cannot be distinguished from that of lower concentrations. Therefore, we determine the sensitivity to be 104 copies/μL. As the plasmid concentration decreases, the Ct value increases (Figure 2B and 2D), which meets the expected results.
Blood Samples Experiments
Cycle 1: Agarose Gel Electrophoresis for Amplification Proof Design
Design
To verify that the genes in the blood sample direct amplification are identical to those in the plasmid experiment, we designed an agarose gel validation experiment. We did a gel test of the blood sample and the plasmid, and determined whether the genes in the blood sample were identical to those in the plasmid by observing whether the band lengths were consistent.
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(1) Prepare 2% agarose gel
1. Weigh 1 g agarose into a conical flask. Add 50 mL TAE buffer. Microwave until boiling. Remove, shake tube, and microwave again until boiling. Repeat this process at least three times until agarose is completely dissolved and the solution is clear and transparent.
2. Allow to cool to room temperature until approximately 50°C. Add 5 µL of 10,000×4S Gel Red nucleic acid dye while gently shaking the flask to ensure uniform dissolution in the buffer.
3. Pour the buffer into a pre-cleaned mold, insert a comb, and allow to solidify (approximately 30 minutes).
(2) Gel Electrophoresis
1. Sample Preparation
Typically, PCR products are mixed with loading buffer before electrophoresis.
However, our PCR Master Mix solution contains integrated loading buffer, allowing direct electrophoresis of the reaction products.
2. Remove the comb from the solidified agarose gel and place the gel into an electrophoresis chamber filled with 1× TAE buffer.
3. Add our product (10 µL) and DNA marker (5 µL) sequentially into the tubes of the agarose gel.
4. Apply power at 150 V for 30 minutes of agarose gel electrophoresis.
5. After electrophoresis, place the gel in a gel viewer to observe the bands.
1. Weigh 1 g agarose into a conical flask. Add 50 mL TAE buffer. Microwave until boiling. Remove, shake tube, and microwave again until boiling. Repeat this process at least three times until agarose is completely dissolved and the solution is clear and transparent.
2. Allow to cool to room temperature until approximately 50°C. Add 5 µL of 10,000×4S Gel Red nucleic acid dye while gently shaking the flask to ensure uniform dissolution in the buffer.
3. Pour the buffer into a pre-cleaned mold, insert a comb, and allow to solidify (approximately 30 minutes).
(2) Gel Electrophoresis
1. Sample Preparation
Typically, PCR products are mixed with loading buffer before electrophoresis.
However, our PCR Master Mix solution contains integrated loading buffer, allowing direct electrophoresis of the reaction products.
2. Remove the comb from the solidified agarose gel and place the gel into an electrophoresis chamber filled with 1× TAE buffer.
3. Add our product (10 µL) and DNA marker (5 µL) sequentially into the tubes of the agarose gel.
4. Apply power at 150 V for 30 minutes of agarose gel electrophoresis.
5. After electrophoresis, place the gel in a gel viewer to observe the bands.
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Gel images of blood and plasmid amplification.
Figure 18. Gel images of blood and plasmid amplification.
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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.
Cycle 2: Two-step amplification method
Design
To investigate the optimal PCR amplification process. We conducted single-round and double-round PCR.
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Using blood samples heterozygous blood samples and wild-type blood samples
1. PCR Reaction System (20 µL):
1. PCR Reaction System (20 µL):
Table 15. The PCR reaction system.
94℃,5min + ( 94℃,30s + 55℃,30s + 68℃,25s) ×34 cycles + 68℃,10min
2. qPCR system(25 µL)：
Table 16. The qPCR reaction system.
Reaction procedure: 94℃,5min + ( 94℃,30s + 55℃,30s + 68℃,25s) ×40 cycles
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Figure 19. 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.
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As shown in the Figure 19, 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.
Cycle 3: Direct Blood Sample Amplification
Design
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.
Figure 20. Flowchart of blood sample experiment.
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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. The reaction system is shown in the table below.
Table 17. The qPCR reaction system for blood in one-step.
94℃,5min + ( 94℃,30s + 55℃,30s + 68℃,25s) ×45 cycles
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Figure 21. Real-Time Fluorescent qPCR. 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.
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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.
Cycle 4: Optimization of Incubation Time
Design
Optimize the incubation time of the red blood cell lysis step to achieve efficient red blood cell removal while maximizing the integrity and detectability of target white blood cell DNA, thereby enhancing the sensitivity and accuracy of subsequent qPCR assays.
Strictly adhere to the single-variable principle. Systematically vary incubation time while maintaining complete consistency in all other lysis conditions (lysis buffer volume, sample volume, temperature (4 °C), mixing method, centrifugation parameters) and subsequent qPCR reaction systems.
Variable Setup: Establish three distinct incubation time gradients: 10 min, 20 min, 30 min.
Control: Perform three technical replicates at each incubation time point to ensure result reliability.
Strictly adhere to the single-variable principle. Systematically vary incubation time while maintaining complete consistency in all other lysis conditions (lysis buffer volume, sample volume, temperature (4 °C), mixing method, centrifugation parameters) and subsequent qPCR reaction systems.
Variable Setup: Establish three distinct incubation time gradients: 10 min, 20 min, 30 min.
Control: Perform three technical replicates at each incubation time point to ensure result reliability.
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1. Sample Preparation: Take 100 μL of heterozygous blood sample.
2. Lysis: Add the sample to a centrifuge tube containing 500 μL of red blood cell lysis buffer.
3. Mixing: Immediately gently invert the tube 10 times to ensure thorough mixing.
4. Incubation: Place the mixture in a 4 °C refrigerator and incubate for 10 min, 20 min, and 30 min, respectively (all ending at the same time).
5. Centrifugation: After incubation, centrifuge the samples at 5,000 rpm for 5 min.
6. Remove supernatant: Carefully discard the supernatant to avoid disturbing the pellet (primarily containing white blood cells and incompletely lysed red blood cells).
7. Resuspend: Add 50 μL of red blood cell lysis buffer to the pellet. Gently but thoroughly pipette or vortex to resuspend the pellet.
8. Template Preparation: Take 5 μL of the resuspension and use it directly as the qPCR reaction template (DNA template).
9. qPCR Reaction Setup: Prepare the qPCR reaction mixture (total volume 25 μL) according to the table below. Ensure the same reaction setup is used for samples from each time point.
2. Lysis: Add the sample to a centrifuge tube containing 500 μL of red blood cell lysis buffer.
3. Mixing: Immediately gently invert the tube 10 times to ensure thorough mixing.
4. Incubation: Place the mixture in a 4 °C refrigerator and incubate for 10 min, 20 min, and 30 min, respectively (all ending at the same time).
5. Centrifugation: After incubation, centrifuge the samples at 5,000 rpm for 5 min.
6. Remove supernatant: Carefully discard the supernatant to avoid disturbing the pellet (primarily containing white blood cells and incompletely lysed red blood cells).
7. Resuspend: Add 50 μL of red blood cell lysis buffer to the pellet. Gently but thoroughly pipette or vortex to resuspend the pellet.
8. Template Preparation: Take 5 μL of the resuspension and use it directly as the qPCR reaction template (DNA template).
9. qPCR Reaction Setup: Prepare the qPCR reaction mixture (total volume 25 μL) according to the table below. Ensure the same reaction setup is used for samples from each time point.
Table 18. qPCR system for Incubation time optimization of blood samples.
Each set of experiments repeated 3 times.
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Place the prepared qPCR reaction plate into the qPCR instrument and run the pre-optimized amplification program (Pre-denaturation: 94°C for 5 min; Cycling: 94°C for 30 sec, 55°C for 30 sec, 68°C for 25 sec, repeated 45 times).
Set the fluorescence channels to dual-channel FAM and ROX.
Set the fluorescence channels to dual-channel FAM and ROX.
Figure 22. 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.
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Red blood cell lysis is a crucial step before DNA extraction and purification from blood samples. It aims to eliminate large quantities of anucleated red blood cells and minimize their interference—such as inhibitors and background DNA—in downstream molecular assays like qPCR. The efficiency of lysis depends on factors including the composition of the lysis buffer, temperature, duration, and sample-to-buffer ratio.
For the specific lysis buffer system and heterogeneous blood samples used in this experiment, the optimal lysis incubation time has not yet been established. Insufficient lysis may leave residual red blood cells, which can increase background noise or inhibit qPCR, while excessive lysis may cause unnecessary damage to target white blood cell DNA or increase the dissolution of impurities.
Lysis incubation time was identified as a key variable affecting lysis efficiency and final qPCR outcomes.
For the specific lysis buffer system and heterogeneous blood samples used in this experiment, the optimal lysis incubation time has not yet been established. Insufficient lysis may leave residual red blood cells, which can increase background noise or inhibit qPCR, while excessive lysis may cause unnecessary damage to target white blood cell DNA or increase the dissolution of impurities.
Lysis incubation time was identified as a key variable affecting lysis efficiency and final qPCR outcomes.
Cycle 5: Optimization of Blood Sample Loading Volume for qPCR
Design
We have established a direct amplification method for detecting folate metabolism genes in blood samples. This approach enables rapid qPCR analysis after blood sample processing and allows identification of folate metabolism-related gene types based on distinct fluorescence results. We aim for widespread adoption in primary care and community hospitals. Therefore, ensuring high efficiency, convenience, and accuracy in direct blood sample amplification is essential, with the goal of achieving precise results using minimal sample volume.
We conducted an engineering cycle to optimize the blood sample loading volume in qPCR. While maintaining all other lysis conditions (lysis buffer volume, sample volume, temperature (4°C), mixing method, centrifugation parameters, incubation time) identical, we designed the blood sample loading volume within the qPCR system. 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.
We conducted an engineering cycle to optimize the blood sample loading volume in qPCR. While maintaining all other lysis conditions (lysis buffer volume, sample volume, temperature (4°C), mixing method, centrifugation parameters, incubation time) identical, we designed the blood sample loading volume within the qPCR system. 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.
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1. Sample Preparation: Pipette 100 μL of heterozygous blood sample.
2. Lysis: Add the sample to a centrifuge tube containing 500 μL of red blood cell lysis buffer.
3. Mixing: Immediately gently invert the tube 10 times to ensure thorough mixing.
4. Incubation: Place the mixture in a 4°C refrigerator and incubate for 20 minutes.
5. Centrifugation: After incubation, centrifuge the sample at 5,000 rpm for 5 minutes.
6. Remove Supernatant: Carefully discard the supernatant, avoiding disturbance of the pellet (primarily containing white blood cells and incompletely lysed red blood cells).
7. Resuspension: Add 50 μL of red blood cell lysis buffer to the pellet. Gently but thoroughly pipette or vortex to resuspend the pellet.
8. Template Preparation: Take 5 μL of the resuspension directly as the qPCR reaction template (DNA template).
9. qPCR Reaction Setup: Prepare the qPCR reaction mixture (total volume 25 μL) according to the table below. Maintain the total volume while designing blood sample additions of 2 μL, 5 μL, and 7 μL. Top up to 25 μL with ddH2O. The blood sample used is sample 3 with a heterozygous genotype.
2. Lysis: Add the sample to a centrifuge tube containing 500 μL of red blood cell lysis buffer.
3. Mixing: Immediately gently invert the tube 10 times to ensure thorough mixing.
4. Incubation: Place the mixture in a 4°C refrigerator and incubate for 20 minutes.
5. Centrifugation: After incubation, centrifuge the sample at 5,000 rpm for 5 minutes.
6. Remove Supernatant: Carefully discard the supernatant, avoiding disturbance of the pellet (primarily containing white blood cells and incompletely lysed red blood cells).
7. Resuspension: Add 50 μL of red blood cell lysis buffer to the pellet. Gently but thoroughly pipette or vortex to resuspend the pellet.
8. Template Preparation: Take 5 μL of the resuspension directly as the qPCR reaction template (DNA template).
9. qPCR Reaction Setup: Prepare the qPCR reaction mixture (total volume 25 μL) according to the table below. Maintain the total volume while designing blood sample additions of 2 μL, 5 μL, and 7 μL. Top up to 25 μL with ddH2O. The blood sample used is sample 3 with a heterozygous genotype.
Table 19. qPCR system for optimization of blood sample loading volume.
Each set of experiments repeated 3 times.
Test
Place the prepared qPCR system into the qPCR instrument and run the pre-optimized amplification program.
Program: 94°C, 5 min + (94°C, 30 s + 55°C, 30 s + 68°C, 25 s) × 40 cycles
Program: 94°C, 5 min + (94°C, 30 s + 55°C, 30 s + 68°C, 25 s) × 40 cycles
Fluorescence channels set to dual FAM and ROX channels.
Figure 23. 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.
Learn
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.
Application of POCT
Design
To detect the genotype of the MTHFR gene using the probe method, we need to amplify DNA from blood samples. While qPCR is straightforward, it requires specialized equipment. Therefore, we attempted to eliminate the multi-cycle temperature control inherent in PCR experiments, which is difficult to achieve under general conditions. We selected Recombinase Polymerase Amplification (RPA) technology. RPA is an isothermal nucleic acid amplification technique. Its core principle involves the binding of recombinase to primers to form a complex. This complex actively scans double-stranded DNA, mediating target sequence recognition and strand invasion. With the synergistic action of single-stranded DNA-binding proteins (SSB), the complex stably unwinds the double-stranded template, preventing re-annealing and thereby replacing the high-temperature denaturation step in traditional PCR. Subsequently, DNA polymerase with strand displacement activity can continuously extend primers at a constant temperature of 37-42°C, achieving exponential nucleic acid amplification. This mechanism eliminates the need for sophisticated equipment like thermal cyclers in RPA reactions. Amplification can be completed within 15-20 minutes under constant temperature conditions, offering advantages of speed, sensitivity, and portability. This aligns with the project's objective of developing rapid detection test strips. Selecting primers with high specificity that successfully amplify the target gene is critical.
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Based on the full sequence of the target gene MTHFR, we designed four primers. The primer sequences are as follows:
Table 20. The different primers for RPA.
Test
To determine which primers successfully amplify the target gene and facilitate experimental result presentation, we performed RPA amplification on wild-type and mutant genes using four primer pairs. The amplification products were visualized via agarose gel electrophoresis.
Figure 24. Proof of amplification of RPA with agarose gel. RPA amplification results for wild-type and mutant genes using four primer sets.
We found that Primer 3 yielded the most effective amplification, though its blank control also produced a band. To rule out aerosol contamination and other interferences, we conducted a separate RPA experiment specifically with Primer 3 to confirm its feasibility.
Figure 25. Proof of amplification of RPA with agarose gel. RPA Amplification Results for Primer 3 Alone.
From the image, we can see that the amplification product size using primer 3 is the expected 185 bp, demonstrating that our designed primer can successfully amplify RPA.
Learn
The agarose gel electrophoresis band pattern aligns with expectations, and the final blank group shows no distinct bands, confirming that our designed primers successfully amplified the target gene. Primers 2 and 4 may exhibit reduced specificity due to shorter fragments, resulting in less pronounced amplification outcomes. Additionally, all four primer sets produced varying degrees of banding in the blank control group. This is likely attributed to the high sensitivity of RPA; further optimization of the RPA system's reaction conditions is required.
Cycle 2: Proof of amplification of RPA with SYBR Green I dye
Design
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.
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To determine the optimal effective concentration of SYBR Green I dye, we designed experiments using 0.5× SYBR Green I. Each reaction system included three experimental groups: wild-type plasmid, mutant plasmid, and blank control.
Table 21. RPA system containing SYBR Green I Dye.
Each set of experiments repeated 3 times.
1. Add Buffer A to the dry powder tube and mix it up and down.
2. Add template (wild mutant), R primer and F primer, mix it up and divide it into two tubes, each of which is 23.125 μL.
3. Add dye and water in corresponding volume respectively, add Buffer B, mix well.
4. Put it into the qPCR instrument for isothermal amplification and fluorescence detection. The program is 38 ℃, 30 min, read the fluorescence once per minute, and the fluorescence channel is set as SYBR Green I.
1. Add Buffer A to the dry powder tube and mix it up and down.
2. Add template (wild mutant), R primer and F primer, mix it up and divide it into two tubes, each of which is 23.125 μL.
3. Add dye and water in corresponding volume respectively, add Buffer B, mix well.
4. Put it into the qPCR instrument for isothermal amplification and fluorescence detection. The program is 38 ℃, 30 min, read the fluorescence once per minute, and the fluorescence channel is set as SYBR Green I.
Test
Figure 26. Real-time fluorescence curve with SYBR Green I.
According to Figure 3, 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.
Learn
The RPA system demonstrated excellent amplification results for samples, with experimental outcomes meeting expectations.
Cycle 3: Proof of the feasibility of the combination of RPA, RNase H II and probes
Design
RPA is an important part of experimental point-of-care testing (POCT), which eliminates the disadvantage of PCR requiring gradient temperature experiments and achieves rapid amplification. We conducted experiments using the RNase H II enzyme and probe from the previous experimental system, combined with RPA, in an attempt to achieve POCT.
To achieve the above design, we first conducted single-tube experiments, placing single-probe in one-tube for isolated reaction to ensure it could accurately recognize its corresponding sequence and produce fluorescence output. Subsequently, we mixed two probes in a single tube for reaction, enabling recognition of the tube's sequence and corresponding fluorescence output. This approach enhances experimental efficiency while minimizing unnecessary interference.
To achieve the above design, we first conducted single-tube experiments, placing single-probe in one-tube for isolated reaction to ensure it could accurately recognize its corresponding sequence and produce fluorescence output. Subsequently, we mixed two probes in a single tube for reaction, enabling recognition of the tube's sequence and corresponding fluorescence output. This approach enhances experimental efficiency while minimizing unnecessary interference.
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We previously obtained fluorescent probes meeting specificity and stability requirements during qPCR experiments. Fluorescent signal was used to evaluate RPA amplification efficiency across samples. The large number of samples amplified via RPA reacting with probes to generate fluorescent signals allowed us to simplify the qPCR's complex temperature cycling requirements. This paves the way for establishing a future detection system that clearly presents results to users without requiring specialized equipment.
Table 22. RPA system containing fluorescent probes.
Each set of experiments repeated 3 times.
1.Take 4 tubes of RPA dry powder and add Buffer A to each tube. Invert the tubes vertically and mix thoroughly.
2.Add the template (wild-type/mutant), probe (wild-type/mutant), R primer, F primer, and RNase H II (control group: equal volume water). Mix well and divide into three equal portions, each containing 18.33μL.
1.Take 4 tubes of RPA dry powder and add Buffer A to each tube. Invert the tubes vertically and mix thoroughly.
2.Add the template (wild-type/mutant), probe (wild-type/mutant), R primer, F primer, and RNase H II (control group: equal volume water). Mix well and divide into three equal portions, each containing 18.33μL.
Table 23. Experimental Classification.
3. Add Buffer B and mix thoroughly
4. Load into the qPCR instrument, set the program, and perform isothermal amplification with fluorescence detection
4. Load into the qPCR instrument, set the program, and perform isothermal amplification with fluorescence detection
Test
To investigate the operating temperature of the probe after the RPA reaction, we conducted three sets of parallel experiments.
Program: 38°C, 30 min → 50°C, 30 min (Simultaneously monitor both FAM and ROX fluorescence channels, with fluorescence measurements taken once per minute.)
Program: 38°C, 30 min → 50°C, 30 min (Simultaneously monitor both FAM and ROX fluorescence channels, with fluorescence measurements taken once per minute.)
Figure 27. Proof of the feasibility of the combination of RPA, RNase H II and probes. Real-time fluorescence amplification curves for 30 min amplification at 38 °C and 30 min amplification at 50 °C. (38 °C for 1 to 30 cycles and 50 °C for 31 to 60 cycles, each cycle lasts 1 min).
Program:38℃, 30min→ 60℃, 30 min (Simultaneously monitor both FAM and ROX fluorescence channels, with fluorescence measurements taken once per minute.)
Figure 28. Proof of the feasibility of the combination of RPA, RNase H II and probes. Real-time fluorescence amplification curves for 30 min amplification at 38 °C and 30 min amplification at 60 °C (38 °C for 1 to 30 cycles and 60 °C for 31 to 60 cycles, each cycle lasts 1 min).
Program: 38 ℃, 30min→ 70 ℃, 30min (Simultaneously monitor both FAM and ROX fluorescence channels, with fluorescence measurements taken once per minute.)
Figure 29. Proof of the feasibility of the combination of RPA, RNase H II and probes. Real-time fluorescence amplification curves for 30 min amplification at 38 °C and 30 min amplification at 70 °C (38 °C for 1 to 30 cycles and 70 °C for 31 to 60 cycles, each cycle lasts 1 min).
(Three parallel sets of each experiment were performed, the data shown are all tie values, and we have not shown the error bars to be able to better present the fluorescence trend)
(Three parallel sets of each experiment were performed, the data shown are all tie values, and we have not shown the error bars to be able to better present the fluorescence trend)
From the three figures above, it can be observed that during the first 30 cycles at 38°C, the fluorescence intensity tends to plateau or slightly increase. When the temperature reaches the set active temperature of RNase H II (cycles 31-60), the fluorescence intensity curve rapidly declines before plateauing or slightly increasing again. Regardless of the reaction template, the fluorescence curves of the positive control group and the plasmid-free blank control group nearly overlap, revealing that the fluorescence signal is unrelated to amplification status. Therefore, we conclude that the feasibility validation for this experiment has failed.
Learn
Probes bind only to single-stranded DNA, whereas isothermal amplification yields double-stranded DNA. At RNase H II temperatures, the double strands remain insufficiently unwound for probe binding, and the probe's binding affinity is not strong enough to compete with complementary strands. Additionally, single-stranded DNA-binding proteins in the RPA system may also hinder probe binding. Further exploration of experimental optimization conditions is required.
Cycle 4: RPA-Probe Temperature exploration
Design
We suspect that the double-stranded DNA amplified by the RPA reaction and the single-strand binding proteins in the reaction system affect probe performance. We decided to apply a high-temperature treatment after the RPA reaction to denature the proteins in the system and separate the double strands while preserving RNase H II activity.
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RNase H II is an enzyme that is resistant to high temperatures, whose activity persists after 90 °C treatment in a short time. Therefore, a 10-minute 90 °C step was introduced between RPA completion and probe reaction initiation.
Test
Program: 38°C, 30 min → 90°C, 10 min → 70°C, 30 min (Simultaneously monitor both FAM and ROX fluorescence channels, with fluorescence measurements taken once per minute.)
Figure 30. RPA-Probe Temperature exploration. Real-time fluorescence amplification curves for 30 min amplification at 38 °C, 10 min amplification at 90 °C and 30 min amplification at 70 °C.
Figure 31. RPA-Probe Temperature exploration. ΔF =Fluorescence intensity at 70th cycle - fluorescence intensity at 40th cycle histogram (At 70°C in the reaction). Error bars: SD, n = 3.
As shown in the Figure 31, a significant difference in fluorescence intensity can be observed between the positive experimental group and the control group.
Learn
In this cycle, we achieved detection of the MTHFR gene genotype with just two temperature changes, laying the groundwork for establishing a simplified detection system. However, the experimentally observed enhanced fluorescence in mutant-type blanks also warrants attention.
Cycle 5: RPA-Probe Temperature exploration
Design
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.
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Dilute RNase H II enzyme concentration tenfold.
Test
Program: 38°C, 30 min → 60°C, 30 min (Simultaneously monitor both FAM and ROX fluorescence channels, with fluorescence measurements taken once per minute.)
Figure 32. 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 33. 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.
Learn
In this cycle, we realized 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.
Cycle 6: Further explore the feasibility of the experiment
Design
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 reaction system separately, verifying the feasibility of two-step method.
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We first configured the RPA reaction system and examined the amplification results after incubating in a metal bath at 38°C for 30 minutes. Then, we added RNase H II and incubated the mixture at 50°C for 60 minutes to achieve separation between the RPA reaction system and the RNase H II reaction system.
Test
Figure 34. Gel images of RPA amplification results
Figure 35. Real-time fluorescence amplification curve of RPA products in RNase H II reaction system. (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.
Learn
Based on all experiments in this cycle, we failed to achieve the goal of accurately screening target genes using RPA. We then reviewed relevant literature to identify the cause. The DNA polymerase I in the RPA kit possesses 5'-3' exonuclease activity. During the amplification phase, single-stranded binding proteins in the RPA system attach probes to the front of the primers. However, DNA polymerase I cleaves these probes bound to the substrate during amplification, prematurely releasing fluorescence. This also caused incompatibility between the RPA system and the RNase H II system. To achieve POCT, the entire system requires modification.
