Nucleic Acid Sample Preparation Methodology
a. Options for analysis of DNA quantity, purity, and quality
Spectrophotometric measurement
Spectrophotometry can be used to estimate DNA or RNA concentration and to analyze the purity of the preparation. Typical wavelengths for measurement are 260 nm and 280 nm. In addition, measurements at 230 nm and 340 nm can provide further information (later for more detail)
A260 readings should be between 0.1 and 1.0 to ensure significance (“Purity analysis”, below). Conversion factors for double-stranded DNA, RNA, and single-stranded DNA are for readings taken at neutral pH (i.e., sample diluted in low-salt, neutral pH buffer). For RNA samples that will be recovered from the cuvette, use RNase-free cuvettes or treat quartz cuvettes to remove RNases. Dilute RNA with buffer that is RNase-free.
Quantitation
Analyzing the UV absorption of a nucleic acid solution at 260 nm provides a simple estimation of the concentration because purines and pyrimidines in nucleic acids show absorption maxima around 260 nm (247–272 nm). Using a 1 cm light path, the extinction coefficient for nucleotides at 260 nm is 20. Based on this extinction coefficient, a 50 μg/mL solution of double-stranded DNA, a 40 μg/mL solution of RNA, and a 33 μg/mL solution (lower for oligonucleotides; see below) of single-stranded DNA all exhibit an absorbance of 1.0 when analyzed in a quartz cuvette in low salt and at neutral pH. Using this property, an estimation of the concentration of the doublestranded DNA, RNA, or single-stranded DNA can be made by inserting these constants (50, 40, and 33, respectively) into the following formula:
Nucleic acid concentration (μg/mL) = A260 × dilution factor × constant
For example, for a double-stranded DNA solution diluted 40× for measurement, with an A260 reading of 0.20, the estimated concentration of the undiluted sample would be 0.20 × 40 × 50 = 0.4 mg/mL or 400 μg/mL.
The estimated concentration of the DNA or RNA preparation can be multiplied by the total sample volume to give the total quantity of DNA/RNA. For example, for an RNA solution of 200 μg/mL in a volume of 100 μL, the total quantity of RNA would be 20 μg.
Note: For oligonucleotides, an A260 of 1.0 represents anywhere from 20 to 33 μg/mL, with the actual conversion factor dependent on the length and base sequence of the oligonucleotide. For a more accurate approximation, consult reference 2.
Purity analysis
Nucleic acids extracted from cells normally require purification to remove protein impurities. The A260/A280 ratio gives an indication of protein contamination; however, this measurement is only an indication and not a definitive assessment. DNA and RNA preparations exhibiting an A260/A280 ratio of 1.7 to 1.9 and ≥ 2.0, respectively, are indicative of samples possessing good purity. Deviations from these values may indicate the presence of potential impurities; however, care must be taken when interpreting spectrophotometric data, and a more suitable indication of purity and quality should also be considered based upon the successful use of the nucleic acid in downstream applications (e.g., restriction enzyme digestion, etc.).
In the absorbance spectrum for nucleic acids, the absorbance at 260 nm is located near the top of a broad peak, whereas the 280 nm reading typically occurs in a steeply sloping region; therefore, small changes in wavelength at 280 nm will result in large changes in absorbance. Consequently, small variations in wavelength accuracy have a much larger effect at 280 nm than at 260 nm. It follows that the A260/A280 ratio is susceptible to this effect, and users are warned that spectrophotometers of different designs may give slightly different ratios.
In practice, the concentration of the nucleic acid sample can also affect the A260/A280 ratio. This is especially evident when a sample exhibits readings that approach the spectrophotometer’s minimum or maximum detection limits. For example, if a solution is too dilute, the 280 nm background reading shows a greater proportional interference and thereby has a disproportionate effect on the final result. Therefore, it is advisable to ensure that the A260 value is > 0.1 but < 1.0 for accurate measurements.
Absorbance values at 230 nm can indicate the presence of additional impurities. This wavelength is near the absorbance maximum of peptide bonds and common buffers such as Tris, EDTA, or chaotropic salts. When measuring RNA samples, the A260/A280 ratio should be > 2.0; a ratio lower than this is generally indicative of contamination with GTC, a reagent commonly used in nucleic acid purification. This reagent absorbs over the 230 to 260 nm wavelength range; therefore, a wavelength scan can be particularly useful when assessing the purity of nucleic acid samples. A correction protocol is often used to compensate for the effects of high background absorbance levels. This is performed at a wavelength distant from those associated with the absorbance of nucleic acids and proteins. The procedure can adjust for the effects of turbidity, particulates, and high-absorbance buffers. NanoVue spectrophotometer from Cytiva possesses a background correction facility that is set at 320 nm. This default correction is automatically applied when measuring the concentration of all nucleic acids. The use of the background correction facility is recommended since spectrophotometric data can be grossly distorted by contaminating particles. This is especially true when analyzing dilute samples.
Agarose gel electrophoresis
The benefit of subjecting nucleic acid samples to gel electrophoresis (in combination with ethidium bromide staining) is that in addition to facilitating the isolation of specific-sized fragments for downstream applications such as cloning, it can also convey information about sample quality, purity, and concentration (quantity).
Quantitation
Electrophoresis can be used to estimate the amount of nucleic acid in a sample by the direct visual comparison to molecular weight markers of known concentration. Experienced molecular biologists routinely make such comparisons; for less experienced investigators there are a number of gel documentation systems available with software that facilitates a quantitative assessment. These assessments are based on comparing the fluorescent intensities of samples with controls or molecular weight markers of known concentrations.
Purity analysis
Electrophoresis can be used to estimate the purity of DNA or RNA preparations. When analyzing genomic DNA or plasmid DNA, the retention of DNA in the wells of electrophoresis gels and excessive DNA smearing often indicates residual protein contamination and nuclease activity, respectively. During genomic or plasmid DNA extractions in the absence of RNase, RNA will remain intact, and the rRNA bands will be visible. The presence of RNA will contribute to an erroneously high A260 absorbance determination, which would result in an over-representation of the concentration of the desired DNA product. During the preparation of total RNA from mammalian cells, two major bands are typically observed after electrophoresis with ethidium bromide. These correlate with the 28S and 18S rRNA, while the majority of mRNA is visualized as a faint smear underlying these two bands. So, for example, enrichment of mRNA using oligo(dT) cellulose would cause a significant reduction in the amount of rRNA present in these samples.
Quality analysis
Electrophoresis of DNA or RNA can provide a general idea regarding the intactness of the nucleic acid of interest. For example, electrophoresis of undigested plasmid DNA provides an opportunity to gauge the performance of the extraction method. In general, the more vigorous the extraction procedure is, the greater the exposure of the plasmid DNA to potential damage. Damage can be assessed by visualization of the amount of intact superhelical plasmid DNA present after purification. Any reduction is indicative of damage. Superhelical plasmid DNA typically runs ahead of covalently closed circular (CCC). In stringent applications such as mammalian cell transfection, extraction methods that facilitate the purification of predominantly superhelical plasmid DNA are generally considered to be more desirable. The size range of genomic DNA up to about 20 kb can be estimated using agarose gel electrophoresis versus a large molecular weight marker. PFGE can be performed to better visualize genomic DNA in excess of 20 kb. RNA integrity can be crudely visualized by comparing the ratio of the 28S and 18S rRNA bands (28S:18S of 2 generally indicates good quality RNA), and analyzing the agarose gel for excessive smearing below the 18S band is indicative of RNA degradation. More accurate quality analysis of RNA can be obtained using the RNA integrity number (RIN; see below). The suppliers of electrophoresis equipment provide detailed information, references, and protocols with their products to address many variants in gel construction and buffer compositions for resolving nucleic acids of different types and sizes. These should be consulted for further details.
RNA integrity number
RNA quality analysis
The RNA integrity number (RIN) is generated by an algorithm designed to estimate the integrity of total RNA. Using RIN, sample integrity is no longer determined by the simple ratio of the ribosomal 28S and 18S bands, but by the entire electrophoretic trace of the RNA sample. This includes the presence or absence of degradation products. In this way, an automatic interpretation and assessment of an RNA sample is generated. The assigned RIN is independent of sample concentration, instrument, and analyst, and therefore represents an unbiased measurement standard for RNA integrity.
RIN values also allow researchers to compare RNA samples, for example, before and after prolonged storage. In addition, RIN ensures repeatability of experiments. For example, if a given RIN value is suitable for a specific experiment, then in general other samples possessing the same value should also be suitable for similar experiments. In another instance, a sample with, for example, a RIN of 5 might not work for microarray experiments but may be appropriate for RT-PCR experiments. The RNA Integrity Database is a repository of user-submitted total RNA traces and is designed to determine the typical total RNA profile for different tissue types, as well as the effects of using different RNA extraction methods and kits. Consult references 3 and 4 for more details.
Phred measurement
DNA quality
Phred quality (q) values express the probability of correctly calling a base-peak in a sequencing electropherogram. They are based on key parameters such as resolution, spectral cross talk, and uniformity of peak spacing. The q value is logarithmically proportional to the probability of correctly calling the identified peak (i.e., a Phred value of 20 is an accuracy of 99%, while Phred30 is 99.9% accurate). The minimal acceptable value for sequencing is a Phred value of 20 (5, 6). Phred values can be used to estimate the quality of any sequenced DNA.
Fluorescent stains
Quantitation
The most commonly used technique for measuring nucleic acid concentration is the determination of absorbance at 260 nm. The major disadvantages of this method are the relative contribution of nucleotides, single-stranded nucleic acids, and proteins to the signal, the interference caused by contaminants commonly found in nucleic acid preparations, the inability to distinguish between DNA and RNA, and the relative insensitivity of the assay (an A260 of 0.1 corresponds to a 5 μg/mL double-stranded DNA solution).
A variety of fluorescent stains are available that circumvent many of these problems. These dyes can be used for both the visualization and quantitation of nucleic acid. These range from the ethidium bromide commonly used for the visualization of nucleic acids during agarose gel electrophoresis to the cyanine dye SYBR™ Green. The latter is routinely used in real-time qPCR to detect the accumulation of PCR products.
Ethidium bromide is a DNA intercalating agent used as a fluorescent nucleic acid stain. When exposed to ultraviolet light, it fluoresces with an orange color, intensifying almost 20-fold after binding to DNA. It is commonly used to detect nucleic acids during gel electrophoresis (e.g., double-stranded DNA derived from PCR amplifications, restriction digests, etc.). It can also be used to detect single-stranded RNA because this molecule usually folds back onto itself, providing local base-pairing for the dye to intercalate. Caution should be used with ethidium bromide because it may be a strong mutagen. It is also widely assumed to be a carcinogen or teratogen, although this has never been carefully tested.
Hoechst (bis-benzimide) dyes are part of a family of sensitive fluorescent stains used for labeling DNA (especially the nuclei) in fluorescence microscopy and for fluorescent-activated cell sorting. Hoechst 33258 is routinely used to quantitate DNA in solution. The dye is essentially selective for double-stranded DNA and does not exhibit any significant interference in fluorescent emission in the presence of proteins or other contaminants. It facilitates the detection and quantitation of DNA concentrations as low as 5 ng/mL.
SYBR Green is generally used for post-electrophoresis staining of double-stranded DNA in agarose or polyacrylamide gels, as well as in real-time qPCR to detect the accumulation of PCR products. The detection limit is ~60 pg per band with 300 nm transillumination (20 pg at 254 nm). SYBR Green can also be used to detect single-stranded DNA and RNA in denaturing agarose/ formaldehyde and polyacrylamide/urea gels, although at a reduced sensitivity. The stain is able to detect as little as 1 to 2 ng of a synthetic 24-mer oligonucleotide on a 5% polyacrylamide gel. This sensitivity is 25 to 50 times greater than can be achieved with ethidium bromide. Due to its exceptional sensitivity, SYBR Green is routinely used in applications when only a limited amount of DNA is available.
SYBR Green is also routinely used in real-time qPCR to detect the accumulation of doublestranded PCR products formed during progressive PCR thermal cycles. When SYBR Green is added to a sample, it immediately binds to all double-stranded DNA present. During PCR, the thermostable DNA polymerase amplifies the target sequence to create PCR products. The SYBR Green then binds to each new copy of double-stranded DNA. As PCR progresses, more PCR products are generated. Since the dye binds to all double-stranded DNA, the result is an increase in fluorescence intensity proportionate to the amount of PCR product produced.
The main advantages of using SYBR Green for quantitative, real-time PCR is that it can be used to monitor the amplification of any double-stranded DNA without the use of a specific probe, thus reducing assay setup and running costs. However the nonspecific nature of SYBR green also represents a major disadvantage; that is, because the dye binds to any double-stranded DNA, it can also bind to nonspecific double-stranded DNA sequences and may generate false-positive signals.
PicoGreen™ is an alternative ultrasensitive fluorescent nucleic acid stain for quantitating doublestranded DNA in molecular biological procedures such as DNA amplification, cDNA synthesis for library production, and DNA fragment purification for subcloning. The PicoGreen reagent exhibits an emission maximum at 530 nm when bound specifically to double-stranded DNA (unbound PicoGreen reagent exhibits minimal fluorescence in solution). The detection range of PicoGreen when bound to double-stranded DNA is 1 to 1000 ng/mL.
RiboGreen™ is a sensitive fluorescent nucleic acid stain for determining the RNA concentration in solutions. The RiboGreen reagent exhibits minimal fluorescence when free in solution. Upon binding RNA, the fluorescence increases more than 1000-fold. However, a disadvantage of the RiboGreen reagent is that it also binds DNA. RNA-DNA mixed samples require pretreatment with DNase to generate an accurate RNA selective assay. The RiboGreen RNA assay is ~200- fold more sensitive than ethidium bromide-based assays and ~1000-fold more sensitive than absorbance measurements at 260 nm.
Regardless of the dye used, the quantitation of a DNA solution is achieved by the direct comparison against a standard curve of control samples of known concentrations. A stock solution of standard double-stranded DNA (e.g., derived from calf thymus or salmon sperm) resuspended in TE buffer is routinely used to generate the standard curve. However, to serve as an effective control it may be preferable to prepare the standard curve with a nucleic acid most similar to the type being assayed (genomic DNA, plasmid DNA etc.). The control should also be treated in a similar way as the experimental samples.
b. Concentration of nucleic acids by precipitation
Concentration of DNA by isopropanol precipitation
If the purified genomic or plasmid DNA is too dilute for the selected downstream application, DNA may be concentrated by isopropanol precipitation.
- Add 0.7 volumes of room temperature isopropanol to the purified sample.
- Vortex, then spin for 15 min at 5,000 × g at 4 °C.
- Remove the supernatant by decanting, taking care not to disturb the pellet.
- Add 2 mL of 70% ethanol that has been pre-chilled to 4 °C. Vortex briefly and spin for 10 min at 5,000 × g at 4 °C.
- Carefully remove the supernatant without disturbing the pellet. Air dry for 5–10 min. Do not overdry the pellet as this will make the DNA difficult to redissolve.
- Resuspend the DNA in the desired volume of a suitable buffer (e.g., TE pH 8.0 or 10 mM Tris- HCl pH 8.5). To ensure the pellet is completely dissolved, incubate at 55 °C for 1 h.
Concentration of mRNA by precipitation
(Adapted from the protocol in QuickPrep mRNA Kit.)
Glycogen solution: 5–10 mg/mL glycogen in RNase-free or DEPC-treated water*
Potassium acetate: 2.5 M potassium acetate (pH 5.0) solution (prepared using RNase-free or DEPCtreated water)
DEPC-treated water: Prepare a 0.1% (v/v) solution of diethyl pyrocarbonate in distilled water, shake vigorously, and allow to stand overnight at room temperature. Autoclave the solution on the following day with the cap loosened. Commercially available RNase-free water may be used instead of DEPC-treated water in this protocol.
* 0.1% DEPC-treated water.
1. Add 75 μL (1/10 volume) of potassium acetate solution and 10 μL of glycogen solution to 0.75 mL of mRNA in RNase-free water. Add 1.5 mL of chilled 95% ethanol (2 to 2.5 volumes) and incubate the sample at -20 °C for a minimum of 15 min. The amount of glycogen should remain constant regardless of sample volume.
2. Collect the mRNA by centrifuging at max. speed in a microcentrifuge at 4 °C for 10 min. If the RNA will not be used immediately, store it in this precipitated state (in ethanol) at -80 °C.
3. Decant the supernatant and invert the tube over a clean paper towel. Gently tap the tube on the towel to facilitate the removal of excess liquid. Wash the pellet carefully by pipetting 1 mL of prechilled 80% ethanol into the tube and gently invert several times. Centrifuge at full speed at 4 °C for 10 min. Decant ethanol and allow to air dry. When all traces of ethanol are gone, redissolve the precipitated RNA in an appropriate volume of RNase-free water. To determine the appropriate resuspension volume, consider the RNA concentration desired, the concentration before precipitation and the volume of the sample subjected to precipitation. However, the percentage of the RNA recovered after precipitation will depend on the total amount present. With 10 μg of RNA, for example, approximately 70% will be recovered. You may therefore wish to redissolve the pellet in a volume 25 to 50% smaller than would be required if all of the RNA were recovered.
c. Estimation of cell density for cultured mammalian cells
Cell numbers should be determined using an automated cell counter (e.g., NucleoCounter™). Alternatively, individual cells should be counted under a microscope using a standard hemocytometer (Hausser Scientific/VWR, #1483). Below is a basic protocol for determining cell numbers using a hemocytometer.
1. Clean a hemocytometer and the short coverslip thoroughly and wipe clean with ethanol.
2. If working with adherent cells, trypsinize the cells and wash once with phosphate-buffered saline (PBS). If working with cells in suspension, pellet the cells at 5,000 rpm for 1 min.
3. Resuspend cells in an appropriate volume of PBS to yield roughly 1 × 106 cell/mL. For example, mammalian cells grown to confluence in a 25 cm2 flask yield approximately 2.5 × 106 cells per flask. If a 25 cm2 flask was used, add 2.5 mL of PBS. Make sure cells are completely resuspended without any visible clumps.
4. Add 10 μL of resuspended cells separately to two chambers of a hemocytometer (under a small coverslip), making sure the solution spreads completely under the coverslip by capillary action.
5. Place the hemocytometer under a light microscope, focus on the cells using lowest magnification and begin counting cells only at the four corner squares and the middle square in both chambers of the hemocytometer grid (7). Count all cells except those touching the middle lines at the bottom and right. Aim to have 50 to 100 cells per square-grid. If the cell count is >150/grid, dilute the cells, clean the hemocytometer, and re-count the cells.
d. RPM calculation from RCF
The appropriate centrifugation speed for a specific rotor can be calculated from the following formula:
| RPM = | 1000 | X √ | (RCF) 1.12r |
Where RCF = relative centrifugal force; r = radius in mm measured from the center of the spindle to the bottom of the rotor bucket; and RPM = revolutions per minute.
For example, if an RCF of 735 × g is required using a rotor with a radius of 73 mm, the corresponding RPM would be 3000.
References
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