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DNA extraction

DNA extraction is the process of isolating deoxyribonucleic acid (DNA) from biological samples, such as cells, tissues, or viruses, by separating it from other cellular components including proteins, lipids, RNA, and debris using physical and chemical methods.^[1] This technique typically involves three main stages: cell lysis to break open the cell membrane and release DNA, removal of contaminants through enzymatic or chemical treatments, and purification via precipitation or adsorption to yield high-quality DNA suitable for analysis.^[2] DNA extraction is a foundational procedure in molecular biology, enabling downstream applications like polymerase chain reaction (PCR) amplification, DNA sequencing, cloning, and forensic analysis.^[3] The importance of DNA extraction lies in its role as the initial step for studying genetic material, facilitating research in genomics, diagnostics, and biotechnology by providing pure DNA free from inhibitors that could interfere with enzymatic reactions.^[1] Historically, early methods relied on manual organic solvents, but advancements have introduced automated, high-throughput techniques to improve yield, purity, and efficiency, particularly for challenging samples like cell-free DNA or microbial communities.^[4] For instance, in metagenomics, effective extraction allows analysis of diverse microbial ecosystems without culturing, revealing functional insights into environmental and health-related microbiomes.^[5] Common DNA extraction methods include organic extraction using phenol-chloroform to partition DNA into an aqueous phase, non-organic approaches like salting-out or proteinase K digestion for protein removal, and solid-phase techniques employing silica columns or magnetic beads for binding and elution of DNA.^[1] Each method is selected based on sample type, required purity, and downstream application; for example, silica-based kits are widely used in laboratories for their speed and consistency, while Chelex extraction suits rapid forensic processing of small samples.^[6] Recent trends emphasize non-destructive methods to preserve specimens for morphological studies alongside genetic analysis, enhancing integrative research in fields like ecology and taxonomy.^[7]

Overview

Definition and Purpose

DNA extraction is the process of isolating deoxyribonucleic acid (DNA) from biological samples, such as cells, tissues, or other materials, by disrupting cellular structures to release the DNA and then separating it from contaminants including proteins, RNA, lipids, and other cellular components.^[1] This isolation ensures the DNA is free from interfering substances that could affect subsequent analyses, typically involving mechanical, chemical, or enzymatic methods to achieve effective separation.^[8] The primary purpose of DNA extraction is to obtain high-quality, purified DNA suitable for a range of downstream molecular biology applications, including polymerase chain reaction (PCR) amplification, DNA sequencing, cloning, and genetic analysis.^[9] These applications rely on intact, contaminant-free DNA to ensure accurate results, such as generating amplicons for genotyping or preparing libraries for next-generation sequencing.^[10] Historically, DNA extraction originated in the late 19th century when Swiss biochemist Friedrich Miescher first isolated DNA, then called "nuclein," from white blood cells in 1869 using basic precipitation techniques with acids and salts.^[3] Methods evolved significantly in the mid-20th century with the development of more refined chemical and chromatographic approaches, enabling higher yields and purity for advancing genetic research.^[11] Successful DNA extraction yields purified DNA quantified in micrograms per sample, with typical outputs varying by source material but often ranging from nanograms to milligrams depending on the starting amount and method efficiency.^[12] Purity is commonly assessed spectrophotometrically using the A260/A280 absorbance ratio, where a value of approximately 1.8 indicates high-quality DNA with minimal protein contamination.^[13]

Applications in Research and Industry

In research, DNA extraction serves as a foundational step for genetic mapping, enabling scientists to construct genome maps by analyzing gene arrangements and regulatory elements across species. For instance, high-quality genomic DNA isolates facilitate whole-genome sequencing to identify genetic variations essential for understanding species evolution.^[14] In evolutionary studies, extraction from ancient samples like bones and teeth has revolutionized the field by allowing reconstruction of mitochondrial and nuclear genomes from extinct species, such as Neanderthals and mammoths, to trace population histories and genetic erosion over time.^[15] Forensic analysis relies on DNA extraction from diverse evidence like blood, saliva, and bone to generate profiles for criminal identification, paternity testing, and disaster victim recovery, with methods optimized for degraded samples to achieve reliable short tandem repeat (STR) profiling.^[16] Gene editing technologies, particularly CRISPR-Cas9, depend on extracted DNA for target validation and functional studies, where purified genomic material from edited cells confirms precise modifications in applications spanning agriculture, medicine, and basic research.^[17] This process supports the design of guide RNAs and assessment of off-target effects, accelerating discoveries in functional genomics. In medical contexts, DNA extraction underpins diagnostic testing for genetic disorders through screening programs that analyze blood spots or buccal swabs to detect conditions like cystic fibrosis in newborns.^[18] For personalized medicine, it enables pharmacogenomic profiling to predict drug responses, such as identifying variants in tumor DNA that guide targeted cancer therapies like those for BRCA-mutated breast cancer.^[18] Pathogen identification in clinical samples benefits from optimized extraction protocols that isolate microbial DNA from infected tissues, improving detection sensitivity in metagenomic sequencing for diseases like sepsis.^[19] Industrial applications leverage DNA extraction in biotechnology for developing genetically modified organisms (GMOs), where extracted plant or animal DNA informs transgenic engineering to enhance traits like insect resistance in cotton and maize.^[20] In pharmaceutical research and development, it supports gene expression analysis to produce therapeutic proteins, such as insulin, by elucidating stress-response pathways in microbial hosts.^[20] Agricultural breeding programs utilize extracted DNA for molecular marker-assisted selection, shortening breeding cycles for traits like rice blast resistance through genomic evaluation.^[20] As of 2025, emerging trends emphasize integration with next-generation sequencing (NGS), where high-quality DNA extracts are crucial for long-read technologies like PacBio to minimize bias in whole-genome applications.^[21] Single-cell genomics demands ultra-pure isolates for low-input sequencing, driving innovations in automated extraction to support precision medicine and oncology research amid rising data volumes.^[21]

Biological Foundations

DNA Location in Cells

In prokaryotic cells, such as bacteria and archaea, the genetic material is primarily housed in a distinct region of the cytoplasm known as the nucleoid, which lacks a surrounding membrane and exhibits minimal compartmentalization compared to eukaryotic cells.^[22] The DNA in these cells typically consists of a single, circular chromosome that is not associated with histone proteins, allowing for a more accessible but still protected genomic structure.^[23] This organization facilitates rapid replication and transcription but presents extraction challenges due to the nucleoid's integration within the dense cytoplasmic matrix.^[24] Eukaryotic cells, found in animals, plants, fungi, and protists, compartmentalize their DNA more extensively, with the majority located in the nucleus as linear molecules organized into chromosomes complexed with histone proteins.^[25] In addition to nuclear DNA, eukaryotic cells contain mitochondrial DNA (mtDNA), which resides in the matrix of mitochondria as small, circular genomes encoding essential respiratory proteins, and in plant cells, chloroplast DNA (cpDNA) performs a similar role in organelles derived from endosymbiotic bacteria.^[26] These multiple DNA locations—nuclear, mitochondrial, and chloroplastic—add layers of complexity to extraction processes, as each is enclosed by double membranes that safeguard the genetic material from environmental damage.^[27] The structural barriers surrounding DNA vary by cell type and contribute to extraction difficulties. Prokaryotic cells, particularly Gram-positive bacteria, feature a thick peptidoglycan layer in their cell walls that encases the plasma membrane, a phospholipid bilayer that maintains cellular integrity.^[28] In contrast, plant cells possess rigid cell walls primarily composed of cellulose microfibrils, providing mechanical support but impeding access to the underlying plasma membrane and nuclear envelope.^[29] Animal cells lack cell walls altogether, relying solely on a flexible lipid bilayer plasma membrane for protection, which simplifies initial access but still requires breaching the nuclear envelope to reach chromosomal DNA.^[30] For instance, in human diploid cells, the nuclear DNA comprises approximately 6.27 billion base pairs, concentrated within the nucleus at a total mass of about 6.41 picograms per cell, highlighting the dense packaging that these barriers preserve.^[31] These cellular architectures collectively pose significant obstacles to DNA release, as detailed in subsequent discussions of extraction barriers.

Barriers to Extraction

Extracting DNA from cells encounters numerous physical barriers that protect the genetic material within. In eukaryotic cells, the plasma membrane serves as the primary lipid bilayer enclosure, while prokaryotic cells feature additional peptidoglycan layers. More rigid structures, such as the cell walls in plants composed of cellulose and hemicellulose, or in fungi made of chitin and β-glucans, significantly impede access to intracellular contents during lysis. These walls provide mechanical strength and must be mechanically or enzymatically disrupted to release DNA. In multicellular organisms, extracellular matrices in animal tissues—consisting of collagen, elastin, and proteoglycans—further complicate extraction from solid samples like muscle or bone, requiring homogenization to break down these fibrous networks. Chemical contaminants pose another major hurdle, as they co-purify with DNA and compromise its integrity or downstream applications. Histone proteins tightly bind to DNA in chromatin, forming nucleosomes that stabilize the genome but necessitate proteolytic digestion to liberate free DNA strands. RNA molecules, abundant in cellular extracts, can contaminate DNA preparations, sometimes comprising 28-52% of the apparent nucleic acid yield and interfering with quantification or amplification. In plant tissues, polysaccharides such as pectin and starch create viscous complexes that trap DNA, while phenolic compounds like tannins oxidize to form quinones that covalently bind and degrade nucleic acids. Biological challenges arise from environmental inhibitors and endogenous enzymes that degrade or inhibit DNA recovery. Humic acids, prevalent in soil samples, co-extract with DNA due to their similar charge and size, potently inhibiting polymerase chain reactions by chelating magnesium ions essential for enzymatic activity. In blood-derived samples, heme from hemoglobin acts as a strong inhibitor, binding to DNA polymerase and reducing amplification efficiency, often requiring specialized removal steps. Additionally, ubiquitous nucleases—enzymes like DNase I—rapidly fragment DNA post-lysis if not inactivated, leading to degradation and quantifiable yield losses; without proper inhibitors or rapid processing, recoverable DNA can decrease by 20-50% due to shearing into non-amplifiable fragments. These barriers highlight the need for tailored cell lysis mechanisms to overcome them effectively.

Core Principles

Cell Lysis Mechanisms

Cell lysis is the critical initial step in DNA extraction, involving the disruption of cellular barriers to release genomic material into a soluble form for subsequent purification. This process targets the plasma membrane, cell wall (in prokaryotes and plants), and nuclear envelope (in eukaryotes), employing biochemical or physical forces to achieve efficient DNA liberation while minimizing degradation. Mechanisms of cell lysis are broadly classified into mechanical, chemical, thermal, and osmotic approaches, often combined for optimal yield depending on sample type.^[32] Mechanical lysis utilizes physical shear forces to break cell walls and membranes, proving particularly effective for tough samples such as Gram-positive bacteria, fungi, and plant tissues with lignified walls. Bead beating, a common technique, involves vigorously agitating cell suspensions with small glass, ceramic, or zirconia beads (typically 0.1–0.5 mm in diameter) in a homogenizer, generating impact and shear that fracture cellular structures. Sonication employs high-frequency ultrasonic waves (20–50 kHz) to produce cavitation bubbles, which collapse and create localized shock waves that shear membranes, though it may generate heat requiring cooling to protect DNA integrity. Grinding, often performed under cryogenic conditions with liquid nitrogen, mechanically pulverizes tissues using mortar and pestle or automated mills, effectively lysing heterogeneous samples like soil or biopsies by direct compression and abrasion. These methods yield high DNA recovery from recalcitrant sources but can produce debris that complicates downstream steps.^[9]^[33]^[34] Chemical lysis disrupts cellular integrity through molecular interactions that solubilize lipids or degrade structural polymers, offering a gentler alternative suitable for sensitive eukaryotic cells. Detergents such as sodium dodecyl sulfate (SDS) insert into lipid bilayers, denaturing membrane proteins and fragmenting the phospholipid structure to form micelles that release intracellular contents. Enzymatic agents complement detergents; lysozyme targets the β-1,4 glycosidic bonds in bacterial peptidoglycan, weakening the cell wall for osmotic rupture, while proteinase K proteolytically digests nucleases and other proteins that might degrade DNA. These agents are typically applied in buffered solutions at mild temperatures (37–56°C) to enhance activity without excessive nucleic acid shearing. Chemical methods excel in scalability for liquid cultures but require careful neutralization to avoid inhibiting enzymatic assays.^[32]^[9]^[34] Thermal lysis induces cell disruption by applying heat to denature membrane-associated proteins and destabilize lipid packing, leading to membrane permeability and content leakage. Temperatures of 70–100°C, often via boiling or controlled heating in a thermocycler, cause rapid expansion and rupture, particularly when combined with chelating agents to inhibit DNases. This approach is simple and reagent-free, making it ideal for point-of-care extractions, though prolonged exposure risks DNA fragmentation. Freeze-thaw cycling, a variant, alternates subzero freezing (forming ice crystals that puncture membranes) with thawing, achieving lysis through repeated mechanical stress from volume changes.^[33]^[34] Osmotic shock exploits differences in solute concentration to induce cell swelling and lysis, primarily effective for cells with permeable membranes like mammalian erythrocytes or osmotically fragile bacteria. Exposure to hypotonic solutions (e.g., distilled water or low-ionic-strength buffers) drives water influx via osmosis, increasing internal pressure until the membrane bursts and releases DNA. This non-invasive method preserves biomolecule integrity better than aggressive mechanical techniques and is often used as a pretreatment for Gram-negative bacteria. However, it is less suitable for walled cells like yeast or plants without adjunct enzymes.^[34]^[32]

DNA Purification Strategies

DNA purification strategies aim to isolate nucleic acids from contaminants such as proteins, lipids, RNA, and salts following cell lysis, ensuring high yield and quality for downstream applications. These methods exploit differences in solubility, density, or binding affinity to separate DNA selectively. Common approaches include solvent extraction, centrifugation-based separation, adsorption techniques, and precipitation, each offering distinct advantages in terms of purity, scalability, and ease of use.^[1] Solvent extraction utilizes organic solvents to partition DNA into the aqueous phase while driving hydrophobic contaminants like proteins and lipids into the organic layer. In this process, an equal volume of a phenol-chloroform-isoamyl alcohol mixture (typically 25:24:1) is added to the lysate, forming an emulsion upon vigorous mixing; upon centrifugation, DNA remains in the upper aqueous phase due to its hydrophilic nature, allowing removal of the interphase containing denatured proteins. This method, a cornerstone of classical DNA isolation, effectively removes a broad range of impurities but requires careful handling to avoid phenol toxicity and potential DNA shearing.^[1]^[35] Centrifugation-based separation leverages density gradients to purify DNA based on buoyant density differences. A prominent example is cesium chloride (CsCl) equilibrium density gradient centrifugation, often combined with ethidium bromide (EtBr) for enhanced resolution of supercoiled plasmid DNA from linear chromosomal DNA. In this technique, the lysate is mixed with CsCl and EtBr, then subjected to ultracentrifugation (typically 40-50 hours at 177,000 × g), forming a gradient where DNA bands at its equilibrium position; EtBr intercalation reduces the density of closed circular DNA, enabling distinct separation. This method yields highly pure DNA suitable for cloning but is labor-intensive and requires specialized equipment. The foundational principles were established in the 1958 Meselson-Stahl experiment for density labeling, later adapted for plasmid purification in 1967. Adsorption methods employ silica-based matrices to selectively bind DNA under specific ionic conditions, followed by elution. DNA adsorbs to silica surfaces in the presence of chaotropic salts (e.g., guanidinium thiocyanate or sodium iodide), which disrupt hydrogen bonding and promote hydrophobic interactions between the DNA phosphate backbone and the silica; contaminants remain in solution. Washing steps remove unbound impurities, and DNA is released using low-salt buffers (e.g., TE buffer) that weaken the binding. This approach, pioneered in 1979, enables rapid, scalable purification without hazardous solvents and is the basis for many commercial spin-column kits, achieving high recovery rates (>80%) for fragments as small as 100 bp.^[36] Precipitation concentrates DNA by reducing its solubility in aqueous solutions using alcohols. Ethanol or isopropanol (2-2.5 volumes) is added to the lysate in the presence of salts (e.g., sodium acetate) to neutralize DNA's negative charge, causing aggregation and pelleting upon centrifugation at low temperatures (-20°C overnight enhances yield). For low-concentration samples (<1 μg/mL), carriers like glycogen (10-20 μg) co-precipitate with DNA, improving visibility and recovery without contaminating downstream reactions. This simple, cost-effective step often follows other purification methods to remove salts and concentrates DNA up to 10-fold.^[37]^[38] Purity of extracted DNA is assessed spectrophotometrically using the OD260/280 ratio, where an ideal value of approximately 1.8 indicates minimal protein contamination, as proteins absorb strongly at 280 nm while nucleic acids peak at 260 nm. Ratios below 1.6 suggest protein or phenol carryover, necessitating re-purification, whereas values above 2.0 may indicate RNA presence. This metric, established through empirical standards, provides a quick proxy for quality before applications like sequencing.^[39]^[40]

Standard Methods

Alkaline Lysis Protocol

The alkaline lysis protocol is a widely used method for isolating plasmid DNA from bacterial cells, exploiting the differential denaturation and renaturation properties of plasmid and chromosomal DNA. Developed in the 1970s by Birnboim and Doly, this technique enables rapid screening of recombinant plasmids and forms the basis for miniprep procedures.^[41] The protocol begins with resuspending a bacterial cell pellet, typically from an overnight culture, in a neutral buffer such as Solution I (50 mM glucose, 25 mM Tris-HCl pH 8.0, 10 mM EDTA), which weakens the cell wall and chelates divalent cations to inhibit nucleases; RNase A is often added to degrade RNA. Next, an equal volume of alkaline detergent Solution II (0.2 N NaOH, 1% SDS) is added and gently mixed, denaturing both chromosomal and plasmid DNA while solubilizing cellular proteins and lipids; the high pH causes chromosomal DNA to denature into single strands that tangle with proteins upon neutralization. The lysate is then neutralized by adding 1.5 volumes of Solution III (3 M potassium acetate, pH 5.5), which protonates the DNA, allowing supercoiled plasmid DNA to renature quickly while chromosomal DNA forms an insoluble precipitate with potassium-SDS complexes and cellular debris. The mixture is centrifuged to pellet the precipitate, and the supernatant containing plasmid DNA is collected for further purification, such as ethanol precipitation.^[42]^[43] This method offers several advantages, including its rapidity—completing in under an hour for minipreps—low cost due to inexpensive reagents, and selectivity for covalently closed circular plasmids, which renature efficiently compared to linear or nicked genomic DNA.^[42]^[41] However, alkaline lysis is not suitable for extracting genomic DNA, as the conditions favor plasmid isolation and may shear larger chromosomal fragments. Additionally, without RNase treatment, there is a risk of RNA contamination in the final preparation, potentially affecting downstream applications like sequencing.^[42]

Phenol-Chloroform Extraction

The phenol-chloroform extraction method is a classic liquid-liquid extraction technique for isolating high-purity genomic DNA from eukaryotic cells and tissues, originally developed in the mid-20th century. It relies on the differential solubility of biomolecules in aqueous and organic phases to separate DNA from proteins, lipids, and other contaminants, yielding intact, high-molecular-weight DNA suitable for downstream applications like cloning and sequencing.^[44] The mechanism involves the addition of a phenol:chloroform:isoamyl alcohol mixture (typically in a 25:24:1 ratio), where phenol denatures proteins by disrupting their hydrophobic interactions and hydrogen bonds, causing them to precipitate or partition into the organic phase.^[45] Chloroform enhances phase separation by increasing the density of the organic layer and aiding in the extraction of lipids and denatured proteins, while isoamyl alcohol reduces foaming during mixing.^[44] DNA, being hydrophilic, remains soluble in the aqueous phase, allowing clean separation after centrifugation.^[45] The procedure begins with cell lysis using a buffer containing detergents (e.g., SDS) and proteases (e.g., proteinase K) to disrupt cellular membranes and digest proteins, typically incubating the sample at 55–65°C for 1–3 hours.^[35] An equal volume of phenol:chloroform:isoamyl alcohol is then added, and the mixture is vortexed vigorously for 10–20 seconds to emulsify phases, followed by centrifugation at 12,000–16,000 × g for 5–15 minutes to form distinct layers: the lower organic phase contains proteins and lipids, the interphase holds debris, and the upper aqueous phase holds DNA.^[35] The aqueous layer is transferred to a new tube, and the extraction may be repeated 1–2 times for higher purity; DNA is then precipitated from the aqueous phase by adding 0.5–1 volume of ammonium acetate or sodium acetate and 2–2.5 volumes of cold ethanol or isopropanol, incubating at –20°C for 1 hour or overnight, centrifuging to pellet the DNA, washing the pellet with 70% ethanol, and resuspending in TE buffer or water.^[35] RNase treatment may be included post-lysis to remove RNA contamination.^[44] Yields from this method are typically high, ranging from 200–400 μg of DNA per 100 mg of mammalian tissue such as liver or kidney, depending on sample type and efficiency of lysis.^[46] The technique produces DNA with A260/A280 ratios of 1.8–2.0, indicating high purity free of protein contamination.^[47] Despite its effectiveness, the method raises significant safety concerns due to the toxicity, volatility, and carcinogenicity of phenol and chloroform, requiring handling in fume hoods and proper waste disposal. By the 2020s, these hazards have driven a shift toward safer, column-based silica kits for routine extractions, though phenol-chloroform remains in use for large-scale preparations and high-molecular-weight DNA isolation where maximum yield and integrity are critical.^[47]

Reagents and Materials

Lysis Agents

Lysis agents are essential chemicals employed in the initial step of DNA extraction to disrupt cellular structures and release genomic material. These agents primarily include detergents, enzymes, and buffers that target cell membranes, walls, peptidoglycan layers, proteins, and nucleic acid contaminants, facilitating efficient cell breakage without excessive DNA shearing.^[48] Detergents such as sodium dodecyl sulfate (SDS) and cetyltrimethylammonium bromide (CTAB) are widely used to solubilize lipid bilayers in cell membranes. SDS, an anionic surfactant, denatures proteins and disrupts non-covalent interactions in lipid membranes, enabling effective lysis across various cell types. Typical concentrations of SDS in lysis buffers range from 1% to 2% (w/v), balancing lysis efficiency with minimal protein contamination.^[49]^[50]^[51]^[52] CTAB, a cationic detergent, is particularly suited for plant tissues due to its ability to complex with polysaccharides and phenolic compounds that otherwise inhibit downstream applications. It lyses plant cell walls and membranes under high-salt conditions, improving DNA yield from recalcitrant samples like leaves and seeds. Standard CTAB protocols incorporate 2% (w/v) in extraction buffers for optimal performance.^[53]^[54]^[55] Enzymatic lysis agents complement detergents by specifically degrading structural components. Lysozyme targets the peptidoglycan in bacterial cell walls, particularly effective for Gram-positive organisms, by hydrolyzing β-1,4 glycosidic bonds. It is typically used at concentrations of 1 mg/mL in lysis buffers for bacterial DNA extraction.^[56] Proteinase K, a broad-spectrum serine protease, digests nucleases and other proteins that bind or degrade DNA, aiding in the complete release of nucleic acids from cellular debris. It is commonly added at 0.1–1 mg/mL during lysis to inactivate endogenous enzymes and enhance yield.^[57]^[58] RNase A is included in some lysis formulations to selectively degrade contaminating RNA, preventing it from co-purifying with DNA and interfering with quantification or amplification. It functions by cleaving phosphodiester bonds in single-stranded RNA, typically at concentrations of 0.1–0.2 mg/mL.^[48]^[1] Buffers like Tris-EDTA (TE) maintain optimal conditions during lysis by stabilizing pH and protecting DNA integrity. Tris acts as a buffering agent to keep the pH around 8.0, while EDTA chelates divalent cations such as Mg²⁺, inhibiting nuclease activity that could degrade DNA. TE is often used at 10 mM Tris and 1 mM EDTA for resuspension post-lysis.^[59]^[60]^[61]

Purification and Precipitation Chemicals

Purification and precipitation chemicals play a crucial role in isolating DNA from contaminants following cell lysis, primarily by removing proteins, lipids, and other impurities while concentrating the nucleic acids. Phenol and chloroform are organic solvents commonly used in the phenol-chloroform extraction method to denature and partition proteins into the organic phase, leaving DNA in the aqueous layer for further processing. This technique, developed as a standard deproteinization step, effectively separates DNA from cellular debris due to phenol's ability to disrupt hydrogen bonds in proteins and chloroform's enhancement of phase separation. Chaotropic agents, such as guanidinium thiocyanate (GITC), are employed in silica-based purification strategies to disrupt hydrogen bonding and facilitate DNA adsorption onto silica matrices by increasing the ionic strength and dehydrating the sample, thereby promoting selective binding of nucleic acids over contaminants. For precipitation, alcohols like ethanol and isopropanol reduce the solubility of DNA in aqueous solutions, allowing it to form an insoluble pellet upon centrifugation. Ethanol is typically used at 100% concentration for initial precipitation (2.5 volumes added to 1 volume of sample, incubated at -20°C for 1 hour) and 70% for subsequent washes to remove residual salts without dissolving the DNA pellet. Isopropanol, requiring only 0.6-0.7 volumes compared to ethanol, offers higher efficiency for precipitating DNA from large sample volumes or low-concentration solutions, as DNA exhibits lower solubility in isopropanol, leading to faster and more compact pellets, though it may co-precipitate more salts. Salts such as sodium acetate aid precipitation by adjusting pH to around 5.2 and providing monovalent cations that neutralize the negative charges on DNA phosphate backbones, enhancing aggregation with alcohols; typically, 0.1-0.3 volumes of 3 M sodium acetate are added prior to alcohol precipitation. After pelleting, DNA is resuspended in TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0), which maintains a stable pH and chelates divalent cations to prevent nuclease activity, ensuring the integrity of the purified DNA for downstream applications.

Method Selection Criteria

Sample Type Considerations

The choice of DNA extraction method is heavily influenced by the biological properties of the sample source, as different materials present unique barriers to cell lysis and contaminant removal. For instance, animal-derived samples often require strategies to handle abundant proteins and clotting factors, while plant tissues demand approaches to counter secondary metabolites and rigid cell walls. Microbial and environmental samples, in turn, involve specific enzymatic or kit-based solutions to address cell wall compositions or inhibitory compounds. These considerations ensure maximal DNA yield and purity while minimizing degradation or contamination. In blood and animal tissues, high protein content poses a significant challenge, necessitating the use of strong proteases such as proteinase K to denature nucleoprotein complexes and facilitate DNA release.^[1] For blood samples, anticoagulants like EDTA are essential to prevent clotting, which could otherwise hinder cell lysis and DNA recovery; EDTA chelates divalent cations required for clotting and inhibits DNase activity without altering DNA quantity.^[62] Tissues, particularly frozen or fixed ones, may require additional mechanical disruption to break down cellular structures, with method selection tailored to tissue type to optimize lysis efficiency.^[1] Plant samples are complicated by polysaccharides and phenolic compounds, which can co-precipitate with DNA or inhibit downstream applications, thus requiring additives like cetyltrimethylammonium bromide (CTAB) to selectively precipitate nucleic acids while removing these contaminants.^[63] Polyvinylpyrrolidone (PVP) is often incorporated alongside CTAB to bind and neutralize phenolics, preventing oxidation and DNA binding.^[64] The tough plant cell walls further necessitate mechanical grinding or homogenization prior to lysis to ensure effective cell disruption.^[63] For microbial samples, bacterial DNA extraction typically involves lysozyme to hydrolyze peptidoglycan in the cell wall, enabling efficient lysis particularly for Gram-positive species with thicker walls.^[65] Viral DNA isolation, by contrast, requires initial lysis of the host cell to access virions, followed by targeted purification to separate viral nucleic acids from host debris.^[66] Environmental samples such as soil or water contain inhibitors like humic acids, which bind to DNA and interfere with extraction and amplification, often demanding specialized kits with inhibitor removal technologies like silica-based binding or proprietary washes.^[67] These kits, such as the DNeasy PowerSoil, incorporate steps to mitigate humic and fulvic acid co-extraction, yielding cleaner DNA suitable for metagenomic analysis.^[68]

Throughput and Cost Factors

Manual methods such as phenol-chloroform extraction represent low-throughput approaches, typically processing one to a few samples at a time with an estimated cost of approximately $0.40–$0.50 per sample due to inexpensive reagents like phenol, chloroform, and ethanol.^[69] These protocols require 2–4 hours per batch, involving multiple manual steps including lysis, organic extraction, and precipitation, which demand skilled labor and basic equipment like centrifuges.^[70] While cost-effective for small-scale research, their labor-intensive nature limits scalability beyond routine laboratory settings. In contrast, high-throughput methods using automated kits or magnetic bead-based systems enable parallel processing of up to 96 samples in 96-well plates, with extraction times reduced to 30–60 minutes per run.^[71] These approaches incur higher costs, around $2–$5 per sample, owing to specialized reagents and consumables like magnetic beads, though bulk purchasing can lower this to under $2 for large volumes.^[72] Automation via robotic workstations minimizes hands-on time, making them ideal for diagnostic applications requiring rapid turnaround. Key factors influencing throughput and cost include labor demands, where manual methods rely on technician expertise versus automated systems that significantly reduce personnel needs; equipment investments, such as centrifuges costing $5,000–$10,000 compared to robotic extractors at $10,000–$30,000; and scalability for high-volume scenarios like COVID-19 diagnostics, where centralized labs processed millions of samples daily using automated platforms to alleviate bottlenecks.^[73]^[74] During the pandemic, such systems facilitated nationwide testing surges by integrating with 96-well formats for efficient workflow scaling.^[75] As of 2025, trends toward cost-optimized kits, including magnetic nanoparticle-based and simplified lysis protocols, have achieved reductions of up to 75% in per-sample expenses compared to standard commercial methods, enhancing accessibility for field and resource-limited applications.^[76] These advancements prioritize eco-friendly, low-reagent formulations while maintaining compatibility with downstream analyses.^[77]

Advanced and Specialized Techniques

High-Molecular-Weight DNA Isolation

High-molecular-weight (HMW) DNA isolation techniques focus on preserving long, intact genomic fragments, often exceeding 100 kilobases up to several megabases, to enable structural analysis without fragmentation. These methods employ low-shear conditions to minimize mechanical damage and enzymatic degradation, ensuring DNA integrity for downstream applications.^[78] A cornerstone method involves embedding cells or nuclei in low-melting-point agarose plugs, which protects DNA during lysis and subsequent manipulations. This approach, originally developed by Schwartz and Cantor in 1984 for separating yeast chromosome-sized DNAs via pulsed-field gradient gel electrophoresis, facilitates gentle in situ lysis to avoid shearing forces that could break large molecules.^[79] Pulsed-field gel electrophoresis (PFGE) preparation using these plugs allows resolution of fragments up to 2 megabases by applying alternating electric fields, as demonstrated in early yeast karyotyping studies.^[79] Gentle lysis buffers, typically containing EDTA to chelate divalent cations and inhibit nucleases, are used alongside proteinase K to digest cellular proteins without introducing nicks.^[80] The protocol generally begins with suspending cells in molten agarose (0.5–1% w/v) and casting into plug molds, followed by solidification and incubation in lysis buffer (e.g., 1% sarkosyl, 1 mg/mL proteinase K, 0.5 M EDTA) at 50–60°C for 1–2 hours to lyse cells in situ. Plugs are then washed extensively in TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) to remove debris and enzymes, often dialyzing against the same buffer overnight. For DNA recovery, plugs can be melted at 65–70°C in the presence of beta-agarase or subjected to electrophoresis to elute intact DNA, yielding fragments suitable for PFGE analysis.^[80] This process has been streamlined for various sample types, achieving yields of e.g., 15–30 μg from 10^9 bacterial cells while maintaining megabase integrity.^[81] Key challenges include preventing nuclease-induced nicks, which can fragment DNA during handling; this is mitigated by nuclease-free reagents and RNase treatment to eliminate RNA that might harbor contaminants. Mechanical shear from pipetting or vortexing is avoided through wide-bore tips and plug-based containment, enabling preservation of ultra-HMW DNA up to 300–400 kb as verified by PFGE.^[82] Yields of megabase-sized fragments are typically lower than for sheared DNA (10–50% recovery), but optimization via extended lysis or chelator enhancements improves outcomes.^[78] These techniques are critical for applications like optical mapping, where HMW DNA is stretched and imaged to generate restriction maps for genome assembly, as shown in early eukaryotic studies resolving fragments over 1 Mb.^[83] In long-read sequencing, such as PacBio HiFi or Oxford Nanopore Technologies platforms, intact HMW DNA supports reads spanning structural variants and repetitive regions, enhancing de novo assembly accuracy over short-read methods.^[78] For instance, agarose plug-derived DNA has enabled telomere-to-telomere assemblies in human genomes by providing scaffolds for complex loci.^[82]

Extraction from Challenging Samples

Extracting DNA from challenging samples, such as those encountered in forensic, ancient, or formalin-fixed paraffin-embedded (FFPE) contexts, requires specialized adaptations to address low quantities, degradation, and contaminants. These samples often yield fragmented or trace amounts of DNA, necessitating methods that maximize recovery while minimizing further damage or external interference.^[84] In forensic applications, touch DNA from surfaces like handled objects presents low-copy number challenges, typically involving fewer than 100 cells and resulting in DNA yields below 1 ng. Silica spin column-based extraction is widely used, as it binds DNA under chaotropic conditions, allowing purification from inhibitors present in trace evidence. The addition of carrier RNA during this process significantly improves yields, often by 4- to 20-fold, in microfluidic or column formats. This approach is particularly effective for low-copy number profiling, enabling short tandem repeat (STR) analysis from minute epithelial cell deposits.^[85]^[86]^[87] For ancient DNA (aDNA), samples from archaeological remains are highly degraded, producing highly fragmented DNA with average lengths of 50–150 base pairs due to postmortem hydrolysis and enzymatic activity. Silica-based extraction methods, developed in the late 1990s, revolutionized aDNA recovery by using spin columns or magnetic particles to isolate these ultrashort fragments (as low as 25-35 bp) from bone, teeth, or sediments under denaturing conditions. To mitigate contamination—a major issue given the low endogenous DNA content (often <1%)—pre-extraction decontamination with dilute bleach (e.g., 3-6% sodium hypochlorite) removes surface contaminants, while UV irradiation of tools and workspaces provides additional protection against modern DNA introduction. These protocols preserve fragile molecules by avoiding harsh chemicals and incorporating UV shielding during handling to prevent photodegradation.^[88]^[89]^[90] FFPE tissues, commonly used in clinical archives, pose challenges from formalin-induced cross-links between proteins and DNA, which fragment nucleic acids and inhibit downstream applications. Reversal of these cross-links is achieved through incubation with proteinase K at 56°C for 1-3 hours, followed by heat treatment at 90-100°C to dissociate methylene bridges, yielding DNA fragments suitable for polymerase chain reaction (PCR) amplification. This enzymatic and thermal combination increases extractable DNA by 2-5 fold compared to untreated samples, though yields remain lower (10-100 ng/mg tissue) due to fixation artifacts.^[91]^[92]^[93] As of 2025, innovations like CRISPR-Cas9-based enrichment have emerged for trace DNA, targeting specific loci to amplify low-abundance sequences in forensic or degraded samples. These methods use guide RNAs to direct Cas9 cleavage and selective capture, enhancing detection sensitivity in low-copy number scenarios by 10-100 fold without broad amplification biases. Such targeted approaches complement traditional extractions, particularly for kinship analysis in challenging traces.^[94]^[95]

Post-Extraction Processes

DNA Detection and Quantification

Following successful DNA extraction, the initial verification involves detecting the presence of DNA and quantifying its concentration to ensure sufficient yield for downstream applications. These methods provide rapid assessments of extracted material, with detection relying on physical, optical, or enzymatic properties of DNA. Spectrophotometry, fluorometry, gel electrophoresis, and quantitative polymerase chain reaction (qPCR) are among the most widely adopted techniques, each offering distinct advantages in sensitivity and specificity.^[96] Spectrophotometry measures DNA concentration through ultraviolet (UV) absorbance, primarily at 260 nm, where nucleic acids exhibit peak absorption due to their purine and pyrimidine bases. Instruments like the NanoDrop microvolume spectrophotometer enable quick analysis of small sample volumes (1-2 μL) without dilution, using a conversion factor where 1 optical density unit (OD) at 260 nm corresponds to 50 ng/μL of double-stranded DNA (dsDNA). This method is convenient for routine lab use but can overestimate concentrations in the presence of contaminants like proteins or phenols, which also absorb at nearby wavelengths.^[97]^[96]^[98] Fluorometry provides higher sensitivity and specificity for dsDNA quantification by using fluorescent dyes that intercalate with double-stranded regions, emitting light only upon binding. The PicoGreen assay, for instance, detects as little as 0.002 ng/μL dsDNA in microplate formats, making it ideal for low-yield extractions, while the Qubit fluorometer achieves detection limits around 0.005 ng/μL using dsDNA-specific dyes that minimize interference from RNA or single-stranded DNA. These assays involve mixing the sample with dye, incubating briefly, and measuring fluorescence with a fluorometer or plate reader, offering accuracy superior to spectrophotometry for trace amounts.^[99]^[100]^[101] Gel electrophoresis separates DNA fragments by size under an electric field in an agarose matrix, allowing visual confirmation of extraction success through band patterns. Staining with ethidium bromide, a DNA-intercalating agent, enables UV light visualization of bands, where fluorescence intensity correlates with DNA quantity and migration distance indicates fragment size when compared to molecular weight markers. This qualitative-to-semi-quantitative approach typically detects 1-5 ng of DNA per band and is essential for verifying intact genomic DNA versus degraded products.^[102]^[103] Quantitative PCR (qPCR) offers functional quantification by amplifying and monitoring specific DNA target sequences in real time, using fluorescent probes or dyes to track amplification efficiency via cycle threshold (Ct) values. By targeting conserved genes like GAPDH or β-actin, qPCR not only measures total extractable DNA but also assesses amplifiability, with sensitivity down to picogram levels per reaction, making it valuable for validating extraction from complex samples. Standard curves generated from known DNA dilutions enable absolute quantification, distinguishing functional DNA from non-amplifiable contaminants.^[104]^[105]

Quality Control Measures

Quality control measures in DNA extraction ensure the isolated nucleic acid is free from contaminants, maintains structural integrity, and performs reliably in downstream applications such as sequencing, PCR, or cloning. These assessments typically involve spectroscopic analysis for purity, electrophoretic methods for integrity, functional assays for usability, and adherence to standardized protocols to identify and mitigate extraction issues. Purity is primarily evaluated using ultraviolet (UV) spectrophotometry to measure absorbance ratios. The A260/A280 ratio assesses protein contamination, with values between 1.8 and 2.0 indicating pure DNA, as proteins absorb strongly at 280 nm.^[39] Deviations below 1.8 suggest protein carryover, while ratios above 2.0 may indicate RNA contamination.^[106] The A260/A230 ratio detects organic contaminants like salts, phenols, or carbohydrates, with ideal values greater than 2.0 for high-purity DNA; lower ratios signal the need for additional purification steps.^[107] These ratios are measured in slightly alkaline buffers (e.g., 10 mM Tris-Cl, pH 7.5) for accuracy, and full spectral scans from 220–320 nm can reveal additional contaminants affecting the 260 nm peak.^[108] DNA integrity, which reflects the absence of shearing or degradation, is assessed through gel electrophoresis and automated capillary electrophoresis. Agarose gel electrophoresis, often with ethidium bromide or safer alternatives like GelRed staining, visualizes DNA as a high-molecular-weight band; smearing indicates fragmentation from mechanical shear or nuclease activity during extraction.^[109] For more precise fragment size distribution, the Agilent 2100 Bioanalyzer employs microfluidics and capillary electrophoresis to generate electropherograms, quantifying the DNA Integrity Number (DIN) for DNA samples or detecting degradation peaks.^[110]^[111] Intact genomic DNA typically shows a predominant band above 20 kb, essential for applications like long-read sequencing.^[112] Functional quality verifies the DNA's suitability for enzymatic reactions, focusing on inhibitor absence and enzymatic accessibility. Restriction enzyme digestion tests involve incubating the DNA with enzymes like EcoRI; complete digestion yielding expected fragment patterns on gels confirms lack of nicks or blocks.^[113] PCR amplification success rates, using universal primers for targets like 18S rRNA, gauge amplification efficiency; failure often points to inhibitors such as heme or humic acids from sample matrices.^[114] Inhibitor detection employs spike-in controls, where known quantities of synthetic DNA templates are added post-extraction; reduced amplification compared to controls quantifies inhibition, guiding dilution or cleanup strategies.^[115] Laboratories adhere to international standards like ISO 20186-2 for specimen handling and processing in genomic DNA examinations, ensuring documented controls for contamination and yield.^[116] The FBI Quality Assurance Standards for Forensic DNA Testing Laboratories mandate reagent blanks and extraction controls to monitor process efficiency.^[117] Troubleshooting low yields or degradation often reveals issues like over-lysis, where excessive mechanical disruption or prolonged incubation activates endogenous nucleases, leading to fragmentation; mitigation includes optimized lysis times and protease addition.^[118] Harsh conditions during lysis, such as vigorous mixing, can also shear DNA, reducing yields—gentler methods like enzymatic digestion are recommended for fragile samples.^[119]

Storage and Preservation

Short-Term Storage Protocols

Following DNA extraction, short-term storage protocols aim to preserve sample integrity for immediate downstream applications, typically spanning days to weeks. For bacterial genomic DNA, purified samples can be stored at 4°C in TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0), which maintains a slightly basic environment to minimize degradation while chelating divalent cations that could promote nuclease activity.^[120]^[121] This condition supports stability for up to several weeks without significant loss of yield or quality, as demonstrated in protocols for bacterial genomic DNA where samples remained viable for short-term use under these parameters.^[120] Repeated freeze-thaw cycles should be avoided, as they can cause shearing and reduce DNA integrity even in buffered solutions.^[122] To optimize concentration and prevent dilution-related issues, DNA is typically eluted in volumes of 50–100 μL during the final purification step, yielding concentrations exceeding 10 ng/μL suitable for applications like PCR or sequencing library preparation.^[123] Lower elution volumes, such as 50 μL, are recommended for samples with limited starting material to achieve this threshold, ensuring sufficient template without compromising recovery efficiency.^[124] Storage containers play a critical role in minimizing loss; low-bind plastic tubes, such as those with proprietary surface treatments, are essential to reduce nonspecific adsorption of DNA to polypropylene walls, which can otherwise lead to significant recovery losses in dilute solutions.^[125] These tubes maintain higher effective concentrations compared to standard vessels, particularly for volumes under 100 μL.^[126] For scenarios involving shipping or temporary transport, DNA pellets can be air-dried at room temperature after ethanol precipitation to remove residual solvents, then rehydrated upon arrival using nuclease-free water to restore solubility without introducing contaminants.^[127] This approach preserves DNA for short-term handling while avoiding refrigeration during transit.^[128]

Long-Term Stability and Best Practices

Long-term storage of extracted DNA requires conditions that minimize hydrolysis, depurination, and enzymatic degradation to preserve integrity for downstream applications such as sequencing or genotyping. Freezing at -20°C or -80°C in ethanol precipitates is a standard method, enabling stability for several years. DNA pellets from ethanol precipitation can be stored directly at these temperatures without resuspension, as the low water activity halts degradative processes; for instance, genomic DNA stored as ethanol precipitates at -80°C maintains usability for PCR and other assays over multi-year periods. Similarly, resuspension in TE buffer followed by freezing at -20°C or -80°C supports long-term archival, with minimal loss observed after 5–10 years in biobanked samples. ^[121]^[129]^[130] Optimal stability during frozen storage depends on maintaining a neutral to slightly basic pH of 7–8 and low ionic strength to reduce acid-catalyzed depurination and metal-ion mediated cleavage. TE buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA) is widely recommended, as the EDTA chelates divalent cations that accelerate degradation, while the low salt concentration (approximately 11 mM total ions) limits electrostatic interactions that could promote strand breaks. Under these conditions at -20°C, DNA degradation rates are exceedingly low, with extrapolated half-lives exceeding 500 years based on temperature-dependent models derived from ancient DNA studies, though practical stability is demonstrated for decades without significant fragmentation. ^[121]^[131]^[132] Lyophilization, or freeze-drying, offers an alternative for ambient-temperature storage by removing water to create a desiccated state that inhibits hydrolytic damage. Nucleic acids preserved via lyophilization with protectants like trehalose retain integrity comparable to frozen controls when stored at room temperature or 4°C for weeks to months, with total DNA yields and amplifiability preserved across taxa such as plants and microbes. This method is particularly useful for field collections or resource-limited settings, as lyophilized samples show no significant degradation after 7 days at ambient conditions, and extended studies indicate viability for years under dry, dark storage. ^[133]^[134] Advanced techniques for archival preservation include DNA banking in biobanks, where standardized protocols ensure traceability and quality over decades. Biobanks employ automated systems for aliquoting and freezing at -80°C, with samples from cohorts like the UK Biobank demonstrating sustained DNA quality for genomic analyses after 10+ years of storage. Emerging methods, such as embedding DNA in silica matrices, enable exceptional longevity; cryosilicification encapsulates whole-blood-derived DNA in amorphous silica, conferring resistance to heat, UV, and oxidative stress, with accelerated aging tests projecting stability for over 1,000 years at room temperature. Similarly, deep silicification enhances penetration for tissue-scale preservation, yielding DNA half-lives estimated at 1,347 years at 20°C in model organisms, while maintaining near-100% genomic fidelity upon retrieval and sequencing. As of 2025, additional room-temperature approaches include polymer encapsulation for biomolecule preservation (CacheDNA) and EDTA-based methods for maintaining DNA integrity in biological specimens during transport and storage. ^[135]^[136]^[137]^[138]^[139]^[140] To ensure archived DNA remains suitable for use, biobanks implement periodic quality control, including spectrophotometric assessment of concentration and purity (A260/A280 ratio ~1.8–2.0) and gel electrophoresis or qPCR to detect fragmentation. Samples are re-evaluated at intervals such as every 5–10 years or prior to release, with metrics like RNA Integrity Number (RIN) analogs for DNA (e.g., DIN) guiding retention decisions; for example, ongoing audits in tissue biobanks confirm <1% failure rates for long-stored DNA in multi-omics workflows. These practices, aligned with ISBER guidelines, mitigate risks from freeze-thaw cycles or container interactions. ^[141]^[136]

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To achieve equivalent accuracy with the Quant-iT PicoGreen assay for dsDNA concentrations below 10 ng/mL, one can either optimize the standard curve analysis ...
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Agarose Gel Electrophoresis for the Separation of DNA Fragments
Apr 20, 2012 · Understand the mechanism by which ethidium bromide allows for the visualization of DNA bands 8. Determine the sizes of separated DNA fragments.
[103]
[PDF] Gel Electrophoresis
DNA to be visible. By comparing to size markers and other controls, one can determine the size or sizes of DNA in a given sample as well as estimate quantity.
[104]
Genomic DNA functions as a universal external standard in ... - NIH
Real-time quantitative PCR (qPCR) is a powerful tool for quantifying specific DNA target sequences. Although determination of relative quantity is widely ...
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A qPCR method for genome editing efficiency determination and ...
Dec 11, 2019 · We developed this real time PCR method based on the sensitivity of Taq DNA polymerase to nucleotide mismatch at primer 3′ end during initiating DNA replication.
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Quantification of DNA - QIAGEN
Pure DNA has an A260/A280 ratio of 1.7–1.9. Scanning the absorbance from 220–320 nm will show whether there are contaminants affecting absorbance at 260 nm.
[107]
[PDF] Assessment of Nucleic Acid Purity - Thermo Fisher Scientific
The 260/230 values for “pure” nucleic acid are often higher than the respective 260/280 values. Expected 260/230 values are commonly in the range of 2.0–2.2.
[108]
DNA sample QC for NGS - QIAGEN
Pure DNA has an A260/ A280 ratio of 1.7–1.9. Note: For accurate A260 /A280 values, measure absorbance in slightly alkaline buffer (e.g., 10 mM Tris•Cl, pH 7.5).Missing: A260/ A280
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How should I QC my genomic DNA samples before sequencing?
Nov 5, 2024 · DNA sample integrity should best be QC-ed by agarose gel-electrophoresis and ethidium bromide staining. "Safe" gel-stains such as Gel-Red work just as well.
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[PDF] DNA, RNA, protein and cell analysis Agilent 2100 bioanalyzer
The Agilent 2100 bioanalyzer was the first commercially available instrument to use microfluidics technology for the analysis of biological samples. Today, it ...
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RNA Quality and RNA Sample Assessment | Thermo Fisher Scientific
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PCR inhibition in qPCR, dPCR and MPS—mechanisms and solutions
Feb 12, 2020 · This review includes mechanisms of specific PCR inhibitors as well as solutions to the inhibition problem in relation to cutting-edge DNA analysis.Missing: restriction digest
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The Wild West of Spike-in Normalization - PMC - NIH
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ISO 20186-2:2019(en), Molecular in vitro diagnostic examinations
This document gives guidelines on the handling, storage, processing and documentation of venous whole blood specimens intended for genomic DNA examination ...
[116]
[PDF] FBI Quality Assurance Standards for Forensic DNA
The standards describe the quality assurance requirements that laboratories performing forensic DNA testing or utilizing the Combined DNA Index System.
[117]
Protocol for minicircle production for gene therapy without ... - NIH
Jul 24, 2025 · CRITICAL: Follow the lysis timing exactly as described. Over lysis will release genomic DNA and contaminate your preparation. Table 2.
[118]
Current Nucleic Acid Extraction Methods and Their Implications to ...
Jul 12, 2017 · The main drawback of this procedure is that it is time-consuming and makes use of toxic chemicals like phenol and chloroform. Moreover, CTAB ...
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Storage of DNA - QIAGEN
Purified DNA should be stored at –20°C or –70°C under slightly basic conditions (eg, Tris·Cl, pH 8.0 or TE buffer; see tables 1 M Tris·Cl and TE buffer)Missing: protocols low- bind
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What's the Best Way to Elute and Store Your Plasmid DNA?
Jun 4, 2020 · Plasmid DNA can be stable at 4°C or even room temperature for a short period, and there are indications that Tris buffer is better than water in ...Missing: volume bind tubes
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Comparison and Optimization of DNA Extraction Methods for Human ...
Mar 4, 2025 · We identified an easy and cost-effective optimized DNA extraction method using Chelex from DBSs, with an elution volume of 50 μL and 1 × 6 mm DBS punch.
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DNA LoBind® Tubes - Eppendorf
In stock 3-day deliveryEppendorf LoBind® Tubes signifcantly reduce sample-to-surface binding to ensure maximal recovery of DNA and RNA molecules. The ideal solution for sample ...
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[PDF] Low DNA-binding properties of Nunc and ABgene storage plates
Oct 17, 2022 · Nunc and ABgene polypropylene storage plates have low DNA-binding characteristics, performing comparably to Eppendorf DNA LoBind plates, and ...
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Plasmid Storage by Filter Paper and Ethanol Precipitation
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Protocols for dry DNA storage and shipment at room temperature - NIH
Jun 24, 2013 · Although trehalose might be sufficient for short-term storage, we recommend Biomatrica and Tris-buffered PVA for long-term storage and for ...
[128]
RNA and DNA Integrity Remain Stable in Frozen Tissue After Long ...
In contrast, many laboratory analyses can be reliably performed on specimens stored at −20 C (Fig. 2) for 2 to 5 years. ...<|separator|>
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[PDF] LONG TERM DNA STORAGE EVALUATION AND METHODOLOGY
Jul 20, 2021 · The ionic strength of the solution also affects depurination rates, so storage in salt solutions—as opposed to a low ionic strength buffer—is.
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Crystal structure of a eukaryotic phosphate transporter - Nature
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Preservation of nucleic acids by freeze-drying for next generation ...
Jan 30, 2017 · Samples were freeze-dried or stored at −80°C as controls. Freeze-dried samples were stored at 4°C or ambient temperature. Investigated ...
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Lyophilized human cells stored at room temperature preserve ... - NIH
Jul 31, 2018 · Our study suggests that freeze-drying of human cells is a suitable alternative for the long-term stabilization of total RNA in whole human cells.
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Assessing the impact of long‐term storage on the quality and ...
The aim of this study was to investigate the long‐term stability of the quality of DNA, RNA, and endocrine markers within blood samples amassed from the biobank ...
[135]
Long-term whole blood DNA preservation by cost-efficient ... - Nature
Oct 21, 2022 · Here we present the method of in situ cryosilicification of whole blood cells, which allows long-term preservation of DNA.
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Deep silicification–assisted long-term preservation of structural and ...
Furthermore, we pioneered cryosilicification technology to achieve the long-term preservation of whole blood DNA for over one thousand years (thermally ...
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Applied Molecular-Based Quality Control of Biobanked Samples for ...
Jul 24, 2023 · This study highlights the significance of quality assurance in biobanking facilities, specifically in the context of high-throughput ...
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UK Biobank Compares DNA Quantification Methods
Apr 26, 2018 · While removing a quantification step can save biobanks time and money, quality assurance and quality control are essential to maintaining high ...