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FAQ

• How to determine OD value of primers?

  1.   Molar Amount and Mass

  1 μg of 1,000 bp DNA = 1.52 pmol

  1 μg of SV40 DNA (5,243 bp) = 0.29 pmol

  1 μg of FX174 DNA (5,386 bp) = 0.28 pmol

  1 μg of lambda phage DNA (48,502 bp) = 0.03 pmol

  1 μg of M13mp18/19 DNA (7,249 bp) = 0.21 pmol

  1 μg of pBR322 DNA (4,361 bp) = 0.35 pmol

  1 μg of pUC18/19 DNA (2,686 bp) = 0.57 pmol

  1 pmol of 1,000 bp DNA = 0.66 μg

  1 pmol of M13mp18/19 DNA (7,249 bp) = 4.78 μg

  1 pmol of SV40 DNA (5,243 bp) = 3.46 μg

  1 pmol of pUC18/19 DNA (2,686 bp) = 1.77 μg

  1 pmol of pBR322 DNA (4,361 bp) = 2.88 μg

  1 pmol of lambda phage DNA (48,502 bp) = 32.01 μg

  1 pmol of FX174 DNA (5,386 bp) = 3.54 μg

  
  

  2.   Absorbance Value and Concentration

  1 OD260 of dsDNA = 50 μg/ml

  1 OD260 of ssDNA = 33 μg/ml

  1 OD260 of ssRNA = 40 μg/ml

  
  

  3.  Molecular Weight

  Average molecular weight of a deoxyribonucleic acid (DNA) base = 333 Daltons

  Average molecular weight of a ribonucleic acid (RNA) base = 340 Daltons

  
  

  4.  Nucleic Acid End Concentration

  For linear DNA: pmol ends = pmol DNA × (number of cuts × 2 + 2) 

  For circular DNA: pmol ends = pmol DNA × number of cuts × 2

  1 μg of linear 1,000 bp DNA = 3.04 pmol ends

  1 μg of linear FX174 DNA (5,386 bp) = 0.56 pmol ends

  1 μg of linear pUC18/19 DNA (2,686 bp) = 1.14 pmol ends

  1 μg of linear SV40 DNA (5,243 bp) = 0.58 pmol ends

  1 μg of linear pBR322 DNA (4,361 bp) = 0.7 pmol ends

  1 μg of linear lambda phage DNA (48,502 bp) = 0.06 pmol ends

  1 μg of linear M13mp18/19 DNA (7,249 bp) = 0.42 pmol ends

   
  

• What is the principle behind DNA sequencing? What are the steps? How accurate is ABI 3730XL sequencing?

  The principle of DNA sequencing is the Sanger method (also known as the chain termination method). The specific steps are as follows:

  
  

  1.  Quantification: Quantify the sample and primers.

  
  

  2.  Sequencing Reaction: Add the quantified template and primers to the reaction system along with reagents like BigDye, and perform the sequencing reaction using a PCR machine.

  
  

  3.  Data Reading: Based on the different fluorescent signals, read the base sequence of the tested sample. ABI (Applied Biosystems) currently claims an accuracy of 98.5% for sequencing results within 800 base pairs using the ABI 3730XL sequencing system.

  
  

• Methods of purification and their detailed explanations

  The synthesized oligonucleotides post-desalting (DSL) are cleaved from the CPG using high-purity ammonia vapor mixed with water vapor under high-temperature and high-pressure conditions. The desalted sequences are then purified via standard-phase chromatography columns. Cartridge purification utilizes reverse-phase purification media, similar to reverse-phase HPLC purification principles. Compared to reverse-phase HPLC, Cartridge purification is more efficient and cost-effective. The reverse-phase purification cartridge usually includes a hydrophobic matrix like C18 silica gel, effectively adsorbing DNA. This allows for easy removal of cleaved protecting groups and shorter primer fragments from the reverse-phase column.

  
  

  PAGE Purification: This method uses denaturing polyacrylamide gel electrophoresis to separate DNA primers, followed by recovery of the target DNA from the gel. PAGE purification is highly effective, providing DNA purity exceeding 90%, particularly useful for purifying long-chain oligo DNA (greater than 50 nucleotides).

  
  

  HPLC Purification: It employs principles of high-performance liquid chromatography (HPLC) to purify DNA primers, especially effective for achieving high purity and sensitivity in analytical or purification tasks. Reverse-phase HPLC, with purity exceeding 90%, is commonly used for analyzing and purifying DNA primers, particularly for modified primers. However, the drawback lies in its higher cost and lower efficiency in bulk production.

  
  

• How to measure the OD value of primers?

  When using a UV spectrophotometer to measure the absorbance of a solution at 260nm for quantification, it's crucial to note the proper usage of the UV spectrophotometer. During measurement, it's advisable to dilute the solution to an absorbance range between 0.2 and 0.8 (extremely high or low absorbance can lead to significant errors).To proceed, thoroughly dissolve DNA powder in a specified volume of water. Take a portion of the solution and dilute it to 1ml. Measure its absorbance in a 1ml standard cuvette; this reading represents the OD value for the measured volume, enabling the calculation of the mother solution's OD value.For instance, upon receiving a tube of DNA powder, dissolve it in 1ml of water to create a mother solution. If 50μL of this mother solution diluted to 1ml yields an absorbance reading of 0.25 in a 1ml standard cuvette, it indicates that this 50μL contains DNA with an OD of 0.25. This implies that the original 1ml mother solution contains DNA of 5OD.

  
  

• Nucleotide Conversion

  1.  Molecular Weight

  MW (Da) = 333 ′ N (number of bases)

  
  

  2.  Concentration

  C (mmol/L or pmol/ml) = OD260 / (0.01 ′ N) C (ng/ml) = OD260 ′ MW / (0.01 ′ N)

  Note: MW —— molecular weight; N —— number of bases; OD260 —— absorbance at 260 nm

  
  

  3.  Melting Temperature of Double-Stranded DNA and Oligonucleotides

  For double-stranded oligonucleotides shorter than 20bp: Tm = 2 (A + T) + 4 (G + C)

  For double-stranded oligonucleotides longer than 20bp: Tm = 81.5 + 16.6 (lg[J+]) + 0.41 (%GC) –(600/N)

  Note: N —— length of the primer (in terms of base pairs); J+ —— monovalent ion concentration

  
  

  4.  Conversion between DNA and Expressed Protein Molecular Weights

  1 kb DNA = 333 amino acids ≈ 3.7 ′ 10^4 Da

  10,000 Da Protein ≈ 270 bp DNA 30,000 Da Protein ≈ 810 bp DNA 50,000 Da Protein ≈ 1350 bp DNA 100,000 Da Protein ≈ 2700 bp DNA

  
  

• Why can't I find my primer sequences in the sequencing report?

  1.  If you've provided a PCR product and can't locate your primers in the sequencing report, don't worry—this is standard. Sequencing works by reading genetic sequences using fluorescence-labeled ddNTPs, but as sequencing primers lack these labels, they won't show up in the results. To obtain your primer sequences, there are two approaches.

  
  

  2.  For shorter PCR products (<800bp), you can sequence using the primer from the opposite end. This allows reading to the sequence's end, giving you the reverse complementary sequence of your primer. However, for longer sequences, a sequencing reaction might not cover the beginning. To tackle this, scientists often clone the PCR product into a suitable vector and sequence it using universal primers on the vector. This provides the complete primer sequence, considering the distance between the universal primer and your insert sequence. Given potential inaccuracies in reading the sequence's start during sequencing, cloning and sequencing offer a more comprehensive view of your PCR product.

  
  

  3.  If your sample is a plasmid or bacterial liquid and the PCR product is cloned into the vector, the cloning direction being random might make finding your primer sequence on one strand challenging. In such cases, try checking the complementary strand.

  
  

  4.  Sometimes, when the sequencing primer is too close to your insert fragment, labs may struggle to find the entire primer sequence. This issue arises due to interference at the sequencing start caused by residual dyes or primer dimers, resulting in a poor starting sequence and potential incomplete reading of your primer sequence.

  
  

• How to dissolve primers?

  Our synthesis report provides the volume of water required to dilute the primer to a concentration of 100 μmoles/L (i.e., 100 pmol/μl) per OD. You can add an appropriate amount of nuclease-free double-distilled water (pH > 6.0) or TE buffer (pH 7.5-8.0) based on your experimental needs. Before opening the cap for dissolution, it's advisable to centrifuge at a speed of 3000-4000 rpm for 1 minute to prevent primer loss upon opening.

• My primers work well in PCR, but I don't get good results for sequencing. Why is that?

  The requirements for primers used in sequencing are higher than those used in PCR reactions. Here are types of primers that are not suitable for sequencing reactions:

  1.  Impure primers: Primers used in sequencing require high purity. Small fragments in the synthesis can directly cause significant background peaks. Hence, primers used in sequencing reactions typically have a sequence length of around 24 bases, as longer primers are harder to ensure purity for.

  
  

  2.  Degenerate, random, or specially labeled primers.

  
  

• How should the synthesized primers be stored?

  The undissolved primers are highly stable and can be stored at -20°C for two years. When dissolved, these primers can be pre-diluted to a concentration of 100 μmol/L (μM) as a stock solution. This stock solution can then be divided into aliquots for storage at -20°C, maintaining stability for eighteen months (repeated freeze-thaw cycles may reduce the shelf life). Users are advised to dilute the concentrated solution to the desired working concentration (either 10 pmol/μl or 20 pmol/μl) before using them in experiments.

  
  

• How to detect the purity of primers?

  An accessible laboratory method involves using PAGE with a specific concentration of polyacrylamide gel containing 7M urea. For primers with <12 bases, a 20% gel is used; for 12-60 bases, a 16% gel is used; and for >60 bases, a 12% gel is employed. Approximately 0.2 OD of the primer is dissolved in a urea-saturated solution or by adding urea powder to the primer solution until saturation. Before loading, denaturation is performed by heating (95°C for 2 minutes).

  
  

  Electrophoresis is conducted at 600V, and after a specific duration (approximately 2-3 hours), the gel is peeled off. Using a fluorescent TLC plate, bands are inspected under a UV lamp. If there are no additional bands below the main band, it indicates high purity. However, occasionally, secondary structure bands may appear above the main band due to incomplete denaturation.

  
  

• Do synthetic primers typically have phosphate groups at the 5' and 3' ends?

  No, both the 5' and 3' ends have -OH groups. If a phosphate group is necessary, please specify it when placing your order. Additional charges will be applied for phosphorylation in such cases.

• What should I do if there are non-specific bands in the PCR reaction with synthetic primers?

  PCR failures can stem from various reasons, considering the following aspects:

  1.  Are the primers and the template properly matched, and what's the homology level?

  
  

  2.  Could the primers themselves possess structural complexities?

  
  

  3.  Are the reagents for the PCR reaction functioning properly?

  
  

  4.  Is the PCR instrument functioning correctly?

  
  

  5.  Are the PCR reaction conditions appropriate? If everything seems normal and the issue persists, we offer the option of re-synthesizing the primers at no additional cost.

• Is codon optimization necessary?

  Essential. Different species favor different codons. Codons preferred in humans might be rare in E. coli. BiOligo provides free codon optimization services using proprietary software for global clients.

• What is the standard free cloning vector of BiOligo?

  (1) pUC series (pUC18/pUC19/pUC57), pBluescript II SK(+), PCR2.1, and similar vectors: contain several commonly used restriction endonuclease recognition sites.

  
  

  (2) pTG19-T vector: suitable for T/A cloning. Additionally, our company offers nearly 300 types of commercial expression vectors (such as pET series, pPIC series) for post-gene synthesis cloning and selection.

  
  

  If specific requirements exist, such as using particular or modified vectors, please provide the relevant information for these vectors to facilitate subsequent experiments promptly.

  
  

• If the A260/A280 ratio is less than 1.8 after setting the OD value for the primers, is the purity of the primers acceptable?

  Due to nucleic acids' strong absorbance near 260nm and proteins' strong absorbance near 280nm, the A260/A280 ratio is commonly used to assess nucleic acid purity extracted from biological samples, typically falling between 1.8 and 2.0. This assessment is based on the assumption that the proportions of A, G, C, and T in the sequence are roughly similar.However, synthesized DNA/RNA differs in this aspect. Synthetic sequences, often short (typically between 20 to 30 bases), exhibit varying proportions of A, G, C, and T, leading to significantly different molar extinction coefficients for each base. Consequently, the A260/A280 ratio of primers composed of different base compositions, especially when sequences have a higher content of C and T bases, may fall well below 1.8. Thus, the A260/A280 ratio cannot reliably determine the purity of synthetic primers.

  
  

• How many PCR reactions can be performed using an annealing buffer of 10mM Tris, pH 7.5 - 8.0, 50mM NaCl, and 1mM of primer with an OD of 13.2?

  In general, a primer of approximately 20 bases with an OD of 2 can be used for around 400 PCR reactions. To prepare the primer, dissolve it in EDTA and mix the desired number of moles for annealing, ensuring the total volume does not exceed 500 microliters. Heat the mixture to 95℃ for 2 minutes, then slowly cool it to room temperature (below 30 degrees Celsius). The annealed product can be stored at 4 degrees Celsius for later use.

• Using a 3% agarose gel electrophoresis to analyze the synthesized primers, multiple bands were observed. Why could this happen?

  Denaturing PAGE electrophoresis is a must for primer analysis. Single-stranded DNA primers easily form complex structures, leading to multiple bands in agarose gel electrophoresis and hindering quantification.

• Can we quantify synthesized primers by measuring band brightness post-electrophoresis with Ethidium Bromide staining?

  Ethidium Bromide stains by intercalating into the double helix of nucleic acids. Synthesized DNA molecules are single-stranded and only exhibit staining when they fold into localized hairpin or partial duplex structures. As different primer sequences vary in their ability to form these structures, their staining capacity differs. Consequently, the quantification of synthesized DNA based on Ethidium Bromide band brightness is unreliable.

• What are the payment terms?

  For new customers, full advance payment is required for the initial three orders. Payment terms for subsequent orders may be adjusted subject to mutual agreement.

  
  

  Accepted payment methods include telegraphic transfer (T/T) and credit card. If you require alternative payment arrangements for your convenience, please contact us to discuss options.

• How many PCR reactions can be done with a primer concentration of 2OD?

  Generally, a primer of approximately 20 bases with a concentration of 2OD can yield at least 400 PCR reactions.

• Will primers degrade when transported at room temperature?

  They won't degrade. Dry primers can be stored at room temperature for at least two weeks without degradation. Since typical shipping times range from 1-3 days, your received primers should not have degraded.

• How should the synthesized fluorescently labeled probe be stored?

  1.  Fluorescent probes must be stored away from light.

  2.  For dry samples, store at -20°C.

  3.  It's highly recommended to dissolve the probe in RNase-free TE buffer (pH 8.0). This results in a more stable probe solution with an extended shelf life. Typically, prepare a stock solution of the probe at 100 pmol/μl, divide it into aliquots (to avoid repeated freeze-thaw cycles), and store at -20°C. Before use, dilute the prepared stock solution to create a working solution (10 pmol/μl or 20 pmol/μl) and store the remaining portion at -20°C.

  
  

• Why do oligo DNA strands of the exact same length not appear in the same position on the gel during electrophoresis?

  1.  Variations in the composition of A, G, C, and T affect their individual electrophoretic velocities.

  
  

  2.  Differences in the three-dimensional structures of DNA strands result in varying migration speeds during electrophoresis. These discrepancies are more noticeable in shorter oligo DNA sequences compared to longer ones.

  
  

• What should be done if sequencing of PCR products post-cloning reveals errors in the primer region?

  Since primer purity is typically not 100%, during clone selection, it's possible to choose clones carrying amplified PCR products from impure primer sequences. In this case, reselecting and sequencing another clone should yield the correct result.

  
  

  If sequencing 2-3 selected clones doesn't show improvement, we'll provide a free primer synthesis redo.
  

  
  

• Why might the plasmid received by the customer be difficult to cut or not fully cleave?

  Possible reasons are as follows:

  1.  Incorrect or absent enzyme recognition sequences on the plasmid.

  2.  Inappropriate conditions during the enzyme cleavage reaction.

  3.  Sensitivity of the restriction enzyme to DNA methylation.

  4.  Incorrect dilution or addition method of the enzyme.

  5.  High concentration of glycerol.

  6.  Partial or complete inactivation of the restriction enzyme.

  7.  Introduction of protective bases causing blockage at the enzyme cleavage sites.

  
  

  
  

  Solutions:

  1.  Check if the plasmid DNA contains DNA sequences recognized by the restriction enzyme.

  2.  Optimize the enzyme reaction system using the provided reaction buffer; increase enzyme concentration or use a fresh batch.

  3.  Check for DNA methylation and the enzyme's sensitivity to it.

  4.  Consider changing protective bases or perform PCR amplification of the target fragment and then enzymatic digestion.

  
  

• Failed protein expression for months; sequencing revealed primer errors. What should be done?

  1.  Always validate the DNA sequence before conducting expression experiments.

  2.  We can offer a free primer synthesis redo.

  3.  In case of a claim, compensation is limited to the product's price range, following international industry standards.

  
  

• How to transport and store plasmids and their bacterial strains?

  We provide two tubes containing plasmids with completely accurate gene sequences (minimum of 3 μg). Plasmids can be transported at room temperature. After dissolution, they should be stored at -20°C and ideally avoid repeated freeze-thaw cycles.

• What are the requirements for sample submission when clients provide samples for services?

  For plasmid samples, they can be provided in the form of stab cultures, glycerol stocks, lyophilized plasmids, or plasmid solutions.

  1.  Stab Cultures: Please provide single colony stab cultures. Stab cultures ensure long-term preservation without a significant decrease in plasmid copy numbers. Place the sample in 1.5ml or 2ml Eppendorf tubes containing solid LB media supplemented with the corresponding antibiotic. Use a sterile toothpick to stab a single colony into the LB agar.

  
  

  2.  Glycerol Stocks: Add overnight grown bacterial culture to sterilized glycerol to achieve a final concentration of 20%. Store it in 1.5ml or 2ml Eppendorf tubes, ensuring proper sealing to avoid contamination.

  
  

  3.  Plasmid: Provide 3-5μg of plasmid dissolved in sterile, deionized water or in a dried form.

  
  

  4.  Note:

  a. Seal the Eppendorf tube containing the sample to prevent contamination or loss during transportation.

  b. Label the plasmid name and resistance on the tube.

  c. Provide additional information about the vector via email or attached note.

  
  

• Why might blunt-ended PCR products be challenging to clone?

  Since the typical PCR primers lack a phosphate group at the 5' end, the resulting PCR products also lack this phosphate group. When attempting cloning into dephosphorylated blunt vectors, they fail to integrate. Moreover, when cloned into non-dephosphorylated blunt vectors, the background noise is significantly high. In such cases, it's advisable to perform phosphorylation (PO4 modification) of the 5' ends of the PCR products.

• Is full-chain S-modification necessary for antisense nucleic acid experiments? What other methods increase nucleic acid stability in organisms?

  S-modification enhances DNA stability, protecting it from nucleases within cells. Fully S-modifying the entire DNA chain does increase stability but lowers its melting temperature (Tm), reducing its efficiency in binding to the target sequence. Therefore, researchers often insert several phosphorothioate bonds (typically 3) at both ends of the DNA fragment. This maintains stability while preserving the antisense DNA's binding capability to the target sequence. However, for injections into live animals, fully S-modifying the entire chain remains more effective in enhancing stability.

• What are the distinctions among FITC, 6-FAM, and 5-FAM labeling?

  They are all derivatives of fluorescein, with 5-FAM and 6-FAM being structural isomers. However, FITC differs from the former two due to its linkage to Oligo (via an isothiocyanate bond) compared to them (which use an amide bond). Although they share the fluorescein chromophore, there is typically no practical distinction in their usage.

• What are the specific distinctions in using dual-labeled fluorescent probes featuring quenching groups TAMRA, Eclipse, or BHQ series dyes?

  Dual-labeled fluorescent probes, incorporating quenching groups like TAMRA, Eclipse, or the BHQ series dyes, are commonly employed as hydrolysis probes, often referred to as 'TaqMan' probes, in Real-Time PCR experiments. These quenching dyes exhibit distinctive characteristics due to their varying spectral properties:

  
  

  1.  TAMRA, functioning as both a fluorescent dye and a quenching moiety, emits fluorescence at higher wavelengths, affecting the detection of the reporter group and leading to relatively higher background fluorescence. Conversely, Eclipse and BHQ series dyes serve as non-fluorescent quenchers, effectively quenching the reporter group without emitting fluorescence themselves. Consequently, probes utilizing Eclipse or BHQ dyes typically exhibit lower background fluorescence, resulting in a higher signal-to-noise ratio and increased detection sensitivity.

  
  

  2.  The efficiency of quenching relies on the spectral overlap between the quenching group and the reporter group. TAMRA exhibits a narrower absorption spectrum, limiting its compatibility with certain reporter groups. Eclipse, with a broader absorption range (390 nm-625 nm), allows quenching of various reporter groups such as FAM, HEX, TAMRA, ROX, etc. Similarly, the BHQ series boasts an even broader absorption spectrum, spanning from 430 nm to near-infrared, enabling quenching of multiple reporter groups like Cy3, Cy5, etc., effectively. Hence, Eclipse or BHQ dyes are preferred in multiplexing PCR due to their versatility in quenching a wide range of reporter groups.

  
  

• What principles should be followed when designing TaqMan probes?

  1.  The probe should be located between the two primers.

  
  

  2.  The probe's GC content should range between 20% to 80%.

  
  

  3.  Avoid long homopolymers, particularly a string of the same base, such as "G."

  
  

  4.  The base "G" should not appear at the 5' end.

  
  

  5.  The probe's Tm (melting temperature) should be 8-10°C higher than the primers', typically ranging from 68 to 70°C.

  
  

  6.  For probes longer than 30 bases, it's preferable to place the quenching group in the middle to prevent high background fluorescence. Additionally, the 3' end of the probe should be modified with a phosphate group to prevent extension during the PCR reaction.

  
  

• What are Molecular Probes? How do they differ in usage compared to common dual-labeled probes like TaqMan probes?

  Molecular Probes are a specific type of dual-labeled probes. They form a hairpin structure by complementing their ends, which brings the reporter and quencher groups close together, resulting in minimal fluorescence. Upon binding to the target sequence, these probes separate, producing a bright signal. Their hairpin structure enhances specificity, making them highly effective in detecting single nucleotide differences (SNPs) compared to standard dual-labeled probes like TaqMan probes. They're commonly used in SNP detection and real-time monitoring of mRNA hybridization, RNA processing, and transcription processes within live cells.