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I am currently studying DNA extraction from various bio-originated samples. I am new to this field, and have learned how commercial DNA extraction kits work for bacteria.
I understand that the very first step is called 'lysis', and that this is supposed to break cells to pull out all the stuff inside them, including DNA.
I am studying cell-free DNA extraction procedure from human blood, and I don't understand why the lysis step is necessary in this context.
If lysis is to break cells, and cell-free DNA is already out of the cell and freely circulating, such a step seems unnecessary.
Is my understanding about lysis right? Then, why is the lysis step necessary during cfDNA isolation?
You want to isolate the cfDNA, and leave anything else in the plasma out of your analysis - essentially, the lysis step is part of the purification.
(I'll elaborate with edits shortly)
Actually, it looks like lysis is not a good idea in cfDNA isolation - lysing cells in your sample will result in contamination by genomic DNA, as discussed here. It would be better to stabilize/fix cells to prevent them from lysing, to avoid this risk.
In other words, your initial instincts were correct!
Why are DNase and lysozyme in lysis steps used during protein purification?
If you are purifying a (often specific) protein, you will need to get rid of as much garbage as you can that might be bound to them.
It depends to an extent on which protein you are after, but generally it is a good idea, especially in preparative purification, to get rid of as many impurities as possible.
As proteins are generally large, and consequentially will appear in the lower bands of your (ultracentrifuged) fraction, you want to get rid of any nucleid acids, especially if the protein you are puryfying concerns the likes of Pol1 , Reverse Transcriptase or any other enzyme involved in Replication/Transcription/Translation..
You can separate the Nucleic acids present in the sample by using a DNAse: this will totally hydrolyse DNA into its constituent parts.
Mind you, there are various DNAse's, all differing in their mechanism and specificity for their substrate. Depending on the DNAse used, it can show either Endo- or Exo- nuclease activity.
More to the point, RNAse is more suitable, as RNA is usually more within the buoyancy/fraction range of proteins, and can be bound to the protein.
Lysozyme : An enzyme that has at least two functions:
It deglycosylates enzymes: Most proteins carried in the bloodstream are Glycosylated: they have a few sugar molecules attached to them. Lysozyme will remove the sugar-residues.
It also has the capacity to break down the cell wall of Gram-Positive bacteria, causing them to lyse . (if you want to purify protein from Gram-positive bacteria)
Any degraded products, either nucleic acid or sugar debris, will appear at the top fractions of the Centrifuge Tube due to their "lighter" buoyancy.
To begin working with DNA we have to first isolate it. The first isolation of DNA was done in 1869 by Friedrich Miescher.  Currently it is a routine procedure in molecular biology or forensic analyses. For the chemical method, there are many different kits used for extraction, and selecting the correct one will save time on kit optimization and extraction procedures.
- Cells which are to be studied need to be collected.
- Breaking the cell membranes open to expose the DNA along with the cytoplasm within (cell lysis).
- Lipids from the cell membrane and the nucleus are broken down with detergents and surfactants.
- Breaking down proteins by adding a protease (optional).
- Breaking down RNA by adding an ribonuclease (optional).
- The solution is treated with a concentrated salt solution (saline) to make debris such as broken proteins, lipids and RNA clump together.
- Centrifugation of the solution, which separates the clumped cellular debris from the DNA.
- DNA purification from detergents, proteins, salts and reagents used during the cell lysis step using various organic solvents (ethanol, isopropanol, phenol chloroform etc.). 
Types of extraction
Organic extraction involves the addition of and incubation in multiple different chemical solutions [7a], including a lysis step, a phenol chloroform extraction, an ethanol precipitation, and washing steps. Organic extraction is often used in laboratories because it is cheap, and it yields large quantities of pure DNA.
Legend: SDS - sodium dodecylsulfate Proteinase K - enzyme PCIA - phenol-chloroform isoamyl alcohol 
Chelex extraction method involves adding the Chelex resin to the sample, boiling the solution, then vortexing and centrifuging it. The cellular materials bind to the Chelex beads, while the DNA is available in the supernatant (upper leyer of solution acfter centrifugating). [7b] The Chelex method is much faster and simpler than organic extraction, and it only requires one tube, which decreases the risk of DNA contamination but it yealds much lower quantity of DNA.
Legend: Chelex resin - ions exchange resin in organic chemistry 
Solid phase extraction such as using a spin-column based extraction method takes advantage of the fact that DNA binds to silica gel. The sample containing DNA is added to a column containing a silica gel or silica beads and chaotropic salts. The chaotropic salts disrupt the hydrogen bonding between strands and facilitate binding of the DNA to silica by causing the nucleic acids to become hydrophobic. This exposes the phosphate residues so they are available for adsorption. [7c] The DNA binds to the silica, while the rest of the solution is washed out using ethanol to remove chaotropic salts and other unnecessary constituents. [7d] The DNA can then be rehydrated with aqueous low salt solutions allowing for elution of the DNA from the beads.
DNA "snot" in cell lysis/protein extraction - (Oct/06/2008 )
There is a sticky gooey snot-like substance in my cell lysate that I am told is DNA. It does not dissolve when boiled in SDS, It won't spin down and I don't know how to get rid of it.
Some have suggested using more extraction buffer (RIPA) but this dilutes the protein too much for my use. Any ideas?
Anything that you can do to shear the DNA will help with the viscosity. Sonication works well, but you can also just pass the sample through a syringe& with a needle >22G.
Bear in mind too, that some cell/tissue samples can also produce a thick 'snot' that is not DNA, but ECM collagen/proteoglycans that can be difficult to solubilize with out very harsh conditions (e.g. >6M Urea or >4M Guanidine).
yes one way is to use more RIPA but your concentration would drop. the best way is to keep on ice all the time, that is why in all protocls say even use cold PBS to wash your cells prior to lysis. your problem is that you didn't do it on ice. keep your cell pellet on ice. and use cold RIPA and immediately put your tube on ice. this should eliminate the sticky clump.
we use DNase to get rid of that clump. just nearly 10ul would do. but why use DNase when you can simply work on ice? don't forget ice. I've seen many people who are not ice friendly. they just don't like keeping their tubes on ice and that's a big mistake.
Anything that you can do to shear the DNA will help with the viscosity. Sonication works well, but you can also just pass the sample through a syringe& with a needle >22G.
Bear in mind too, that some cell/tissue samples can also produce a thick 'snot' that is not DNA, but ECM collagen/proteoglycans that can be difficult to solubilize with out very harsh conditions (e.g. >6M Urea or >4M Guanidine).
Thank you for your advice. I tried both sonication (this is the low power, cleaning type) and passing the sample through a #27 needle and this did decrease the viscosity quite a bit but there was still goo that stuck to the pipette tip. I then passed the sample through a needle with a 5um filter installed and this caught all the goo but I am afraid of loosing DNA-interacting proteins that might be bound up in the goo.
There was additional advice offered that suggested keeping the sample cold which I will try today, then the urea/guanidine as a last resort.
I've had samples that I needed so concentrated that I had similar problems. Re-Boil the sample for a minute or two and load while it's still hot. A hot sample is not nearly as gooey as a room temp sample. The only major issue with samples this concentrated is that some proteins may not be able to enter the gel properly. You may want (if you can) to use lower percentage gels.
The problem with leaving the DNA in the sample is the likelihood of its creating variability between samples e.g. one pipette pulls out a wad of goo and the next does not. It turns out that the keeping everything cold idea did work and allowed me to pellet the DNA. My sample protein concentration is around 6.5 mg/ml which is fine for loading a gel etc. Since this is the simplest of all ideas it must be the best. I might try loading the "DNA" pellet on a gel just to rule out the possibility of loosing DNA-binding proteins but otherwise I will stick with this protocol.
Thanks to all who offered their advice!
Naive question, possibly, but is there any reason you can't just add some DNase I?
Improved, two-stage protein expression and purification via autoinduction of both autolysis and auto DNA/RNA hydrolysis conferred by phage lysozyme and DNA/RNA endonuclease
We report improved release of recombinant proteins in E. coli, which relies on combined cellular autolysis and DNA/RNA autohydrolysis, conferred by the tightly controlled autoinduction of both phage lysozyme and the non specific DNA/RNA endonuclease from S. marcescens. Autoinduction occurs in a two-stage process wherein heterologous protein expression and autolysis enzymes are induced upon entry into stationary phase by phosphate depletion. Cytoplasmic lysozyme and periplasmic endonuclease are kept from inducing lysis until membrane integrity is disrupted. Post cell harvest, the addition of detergent (0.1% Triton-X100) and a single 30 minutes freezer thaw cycle results in > 90% release of protein (GFP). This cellular lysis is accompanied by complete oligonucleotide hydrolysis. The approach has been validated for shake flask cultures, high throughput cultivation in microtiter plates and larger scale stirred-tank bioreactors. This tightly controlled system enables robust growth and resistance to lysis in routine media when cells are propagated and autolysis/hydrolysis genes are only induced upon phosphate depletion.
Results and Discussion
Microbial concentration in the GB sediment sample was 1.96 × 10 10 cells/g as determined by fluorescence microscopy (Table 1). This sample was rich in hydrocarbons (oil-immersed), which were derived from terrestrial organic matter and hydrothermal activity. The hydrothermal chimney sample from EPR was rich in silicate, sulfide and iron minerals, and its cell concentration was 1.69 × 10 9 cells/g. The SCS pelagic sediment sample contained 𢏁.4 wt% of organic matter (Wang et al., 2010), and its cell concentration was 1.17 × 10 9 cells/g. The qPCR quantification of the bacterial and archaeal 16S rRNA gene copy numbers indicated that the GB sample was dominated by archaea while bacteria dominated EPR and SCS samples (Table 1).
The M-SDS method recovered the maximum DNA amount among the three different seafloor samples, which was up to 𢏄 times higher than the other two methods (Table 1). This result was further confirmed by fluorometric method and quantification of the DNA concentration based on the band intensity on the agarose gel (Figure 2). Compared with other two methods, longer fragments of the total DNA were recovered by M-SDS, showing a clear main band on the agarose gel (>23 kb) despite some DNA fragmentations occurred. We should emphasize that the focus of this study is DNA extraction from seafloor samples where relatively high biomass are present. Although, we also tested DNA extraction from low-biomass subsurface sediment samples ( 3-4 cells/g sediment), the DNA quantity obtained by all methods was below quantification limit, therefore no conclusion could be made and not included in the study.
FIGURE 2. (A) Agarose gel electrophoresis of the recovered DNA from the Guaymas Basin sediment sample. Equal amount of sample (0.3 g) were extracted with three methods. The loading volume of the recovered DNA was 1 μL. M indicates Lambda DNA/HindIII Marker 2 (Thermo Fisher Scientific, USA). (B) Representation of DNA band intensity (A) as plotted in three dimensional image. (C) Standard curve for measuring the DNA concentration using CLIQS 1D Pro software as determined by the DNA band intensity on the agarose gel.
The highest yield of DNA was achieved by M-SDS possibly due to the combination of multiple cell lysis treatments, including physical (bead-beating), chemical (SDS surfactant), and enzymatic (proteinase K and lysozyme) steps. During the modification of Zhou’s SDS-based DNA extraction method, the addition of a bead-beating step always got higher DNA yields compared to cell lysis treatments tested in this study (i.e., sonication and freeze-thaw). Particularly, the bead-beating step was carefully modified with different conditions, including the intensity of the tissue lyzer, the size of the glass beads, and the ratio of glass beads to sample, resulting in the best optimized condition (higher DNA yield and longer fragment) of 30 Hz intensity, 0.1 mm diameter glass bead, and 1:1 of the bead to sample ratio (Figure 1). Moreover, a twice of ethanol-wash step was used in the final purification instead of using the column filtration (as used by HA and KIT), which may further reduce the chance of DNA breakdown and loss. Substantial DNA loss may occur during silica column purification due to the competitive binding of co-extracts (such as humic acid) to silica membranes, which was not used for all three methods tested in this study (Howeler et al., 2003 Lloyd et al., 2010 Lever et al., 2015). DNA extracts without the final column purification step by HA and KIT methods could not be amplified by PCR (data not shown). The ethanol-washed DNA could be further used for PCR-amplification and metagenomic sequencing without any additional purification steps.
The environmental DNA extracted by the M-SDS method was characterized by higher yields and longer fragments (Figure 2 Table 1). It was proved suitable for the metagenomic study since a high quality metagenomic sequence assemblage was created from the GB sample (see below for more details). In contrast, highly fragmented DNA were obtained by HA and KIT methods (Figure 2), indicating that DNA was physically and/or chemically broken during the extraction steps. For the HA method, DNA was denatured into single strand and might have lost its structural stability due to the high alkaline condition (Ageno et al., 1969 Kouduka et al., 2012). In addition, the high temperature treatment (70ଌ for 20 min) might damage DNA, making the length shorter than 23 kb as reported previously (Morono et al., 2014). The KIT method also yielded less concentrated DNA with relatively short fragments.
The copy numbers of archaeal and bacterial 16S rRNA genes recovered from GB, EPR, and SCS samples by three DNA extraction methods were determined by qPCR. M-SDS recovered the highest 16S rRNA gene copies both for archaea and bacteria, which was consistent with the highest DNA yield (Table 1). In the GB sample, for example, the M-SDS method obtained 7.19 × 10 9 archaeal 16S rRNA gene copies/g and 9.98 × 10 8 copies/g for bacteria, which were 3𠄵 folds higher than those in DNA extracted by HA, and >10 3 folds higher than those using the commercial kit. Surprisingly, the number of archaeal and bacterial 16S rRNA gene copies obtained by KIT was approximately 2𠄳 orders of magnitude lower than those by M-SDS and HA. This was possibly attributed to its substantial loss of DNA during extraction steps, together with the fragmented DNA, which finally reduced the number of 16S rRNA gene templates for the qPCR analysis.
In addition, a higher archaeal to bacterial 16S rRNA gene copy ratio was observed by the M-SDS method from GB (7.2) and EPR (0.28) samples. Since archaeal cell membrane is generally more rigid than bacteria (Valentine, 2007), they are overall highly resistant to cell lysis for the DNA extraction. Nevertheless, M-SDS had a better performance of recovering archaeal DNA, which would provide less-biased archaeal community data. It has noted that harsher extraction methods sheared the bacterial DNA (Zhou et al., 1996 de Lipthay et al., 2004), which may cause the higher ratios of archaea vs. bacteria in the samples. However, this would not be the case here, as the M-SDS method recovered long, less-sheered DNA. This is important for microbial communities inhabiting various seafloor environments, where archaea tend to play a significant role in energy metabolisms and elemental cycles (Lloyd et al., 2013 Meng et al., 2014 Vigneron et al., 2014).
Cell Lysis Efficiency
Cell lysis efficiency was determined by comparing cell concentrations in the sample before and after DNA extraction. In general, between 82.9 and 99.0% of lysis efficiencies were obtained by using three methods examined, although different samples resulted in varied lysis efficiencies (Table 1). In the GB sample, HA and M-SDS lysed 98.6 and 98.2% of the total microbial cells, respectively. These results were slightly higher than that with the KIT (92.5%). The highest cell lysis efficiency was observed in the HA method (98.6%), indicating the advantage of hot-alkaline incubation for archaea-dominated samples or deep subseafloor sediments as reported previously (Morono et al., 2014). For the EPR chimney sample, however, the cell lysis efficiency by the HA method was 82.9%, although KIT and M-SDS methods gave the better performance of cell lysis efficiency, which were 98.8 and 97.4%, respectively. This may be likely due to the dissolution of high metal contents in the chimney sample (James et al., 2014), thus changing the extraction buffer pH or inactivating the lysing enzymes. For example, (1) high concentrations of iron and manganese ions may form hydroxide precipitates and lower the buffer pH (2) heavy metals may cause the inactivation of proteinase K and lysozyme. For the SCS sediment sample, the M-SDS method gave the best performance of cell lysis efficiency (99.0%), followed by HA (96.3%) and KIT (94.9%). The SCS sample was characterized by abundant clay particles (Liu et al., 2010), which may adsorb a significant amount of the extracted DNA (Barton et al., 2006). The high cell lysis efficiency obtained by M-SDS was possibly due to the extraction buffer contained more anionic and cationic surfactants, which might help to isolate the adsorbed DNA from the charged clay minerals (Dias et al., 2004, 2008 Rosa et al., 2005).
Comparison of Microbial Communities
The highest number of archaeal and bacterial 16S rRNA gene sequences were obtained by M-SDS, followed by HA and KIT, although the equal amount of PCR-amplified DNA were sequenced in the same run (Table 2). The reason is not clear yet, but it was suggested that higher quality of DNA helps to increase the target amplification percentage and reduces the chances of unspecific amplification and chimera formation, thus producing more qualified amplicons for sequencing (Martin-Laurent et al., 2001 Scupham et al., 2007 Sergeant et al., 2012).
The rarefaction curve of bacterial communities indicated that most bacterial taxa have been covered at this sequencing depth, although the SCS sediment sample did not plateau (Figure 3). However, the archaeal rarefaction curve gave varied results. As expected, the microbial community structure determined by three DNA extraction methods shared much similarity in each sample. In the GB sediment sample, Bathyarchaeota (formerly referred as Miscellaneous Crenarchaeota Group [MCG]) and ANMEs were predominant in the archaeal community, followed by Thermoplasmata and Parvarchaea. Compared to other two methods, the KIT method recovered higher ratios of Bathyarchaeota (64%) and lower ratios of ANMEs (26% Figure 4). For the bacterial community, Proteobacteria-related sequences were also enriched from the KIT method. The M-SDS and HA methods resulted in similar microbial communities, showing predominance of ANME-1, Thermoplasmata for Archaea, and Candidatus microgenomatus and Firmicutes for Bacteria. It is reported that ANME groups often occurred as the aggregate which are covered by thick extracellular polymeric substances as well as carbonates and encrusted minerals (Boetius et al., 2000 Knittel and Boetius, 2009 Chen et al., 2014). The combination of bead-beating, SDS surfactant, and enzymatic lysis steps in the M-SDS protocol helps to (1) remove cells from the aggregated consortium with minerals (Miller et al., 1999 Niemi et al., 2001 Luna et al., 2006 Morono et al., 2014), and (2) break the rigid cell wall of gram-positive bacteria, likely resulting in higher ratios of ANME- and Firmicutes-related sequences. The similar trend was also seen from EPR and SCS samples. These observations suggest that M-SDS and HA methods may have a superior performance of isolating some archaeal or aggregated microbial communities. For the EPR sample, Parvarchaeota (48.0.1%) and Thermoplasmata (16.5.2%) dominated the archaeal community, whereas Proteobacteria (20.9.9%) and Bacteroidetes (14.8.1%) dominated the bacterial community. For the SCS sample, major groups are Parvarchaeota (29.5.2%), Thaumarchaeota (32.5.2%), followed by minor groups such as Bathyarchaeota (4.2.5%) and MHVG (5.1𠄸.3%), whereas bacterial communities were dominated by Proteobacteria (55.0.18%) and Bacteroidetes (19.1.6%).
FIGURE 3. Rarefaction curves for (A) archaeal and (B) bacterial 16S rRNA gene sequences obtained from three seafloor samples by three different DNA extraction methods. Sequences were clustered into OTUs at 97% similarity cutoff. X and Y axis are not presented on the same scale.
The laboratory equipment used in this class includes the following: powdered soybeans (transgenic and non-transgenic) (Soy Bean Powder SB-Set Fluka), gloves, Eppendorf tubes (standard and PCR-type), ice boxes, a balance, a heating bath, vortexes, centrifuges, several sets of pipetman (adjustable volume: P10, P200, and P1000) with tips, a DNA plant extraction kit (DNeasy™ plant kit), a PCR kit (Taq PCR core kit) from Qiagen, synthetic oligonucleotides (Isoprim), PCR Eppendorf thermocyclers (PolyLabo), agarose electrophoresis systems with generators (Amersham Biosciences, Inc.), agarose (Euromedex), 10 × TBE buffer (Euromedex), a 100-bp DNA ladder (Promega), a microwave to melt the agarose, concentrated ethidium bromide (EB) solution (Sigma), a plate to stock the EB, a spatula to move the gels in and out of the EB bath, an EB-dedicated dustbin, a UV table, and a system to photograph the gels.
The students are encouraged to read the guides for the plant DNA extraction kit (DNeasy™ plant kit) and for the PCR kit (Taq PCR core kit) from Qiagen (France) provided before the practical experiment. They are also given the address of a comprehensive web site, ohioline.ag.ohio-state.edu/gmo/, which is maintained by scientists from the College of Food, Agricultural, and Environmental Sciences at the Ohio State University. This site is organized as a web link to extensive and unbiased GMO information. Thus the students can visit the sites of industrials such as Monsanto and Novartis but also of opposition campaigners such as Greenpeace. This individual homework should lead to a good understanding of the principles underlying the method to be used. During the practical class itself, the students should use the first 2 h to set up a protocol and to explain to the teacher all the steps to be performed. The rationale for each step should be fully understood.
The students, working in pairs, are then ready to perform a 6-h experiment on two samples, one of transgenic and the other of non-transgenic soy flour, according to the detailed instructions given below. It includes the use of the kits to extract the DNA and amplify it by PCR, as well as the analysis of the results by electrophoresis.
Extraction and Purification of Plant DNA—
The isolation of DNA from soybean with the DNeasy™ plant kit is carried out according to the following instructions and by using AP1, AP2, AP3, AW, and AE solutions and the two types of column provided with the kits by the manufacturer (Qiagen). Briefly, the instructions consist of the following: 1) Switch on the heater at 65 °C to keep the AE buffer hot. 2) Weigh 0.1 g of soy powder in a 2-ml Eppendorf tube (maximum capacity of the kit). 3) Add 400 μl of AP1 buffer (the lysis buffer is constituted of detergents, proteases, and salts) and 4 μl of RNase (100 mg/ml) (to avoid the purification of the RNA and permit the liquefaction of the solution), and mix gently. Solution AP1 contains detergents, so avoid contact with skin. 4) Incubate 10 to 15 min at 65 °C (to keep endogenous DNases inactive, denature the proteins, and break the cell walls). 5) Add 130 μl of AP2 buffer, vortex the solution, and put it on ice for 5 min (to precipitate the proteins and the polysaccharides by acidification of the medium). Ice permits the precipitation of unfolded proteins, causes misfolding of other ones, and inactivates the DNases. Solution AP2 contains acetic acid, so avoid contact with skin (material safety data sheet number 64-19-7). 6) Centrifuge the mix for 2 min at 12000 × g on a QIAshredder spin column (to eliminate the cell walls and the precipitate). This column is a filter. It stops the particles only in function of their size. All the soluble molecules can pass through it. 8) The solution is transferred to a fresh 2-ml Eppendorf tube, and the pellet is discarded. By this way, they can evaluate the volume of solution they have and thus calculate how much solution they will add afterward. At this step it is useful to show them how to measure a volume using a pipetman. 9) Add 0.5 volume of AP3 solution, and vortex the mix (to constitute the DNA salts). To avoid chloride production, do not mix AP3 solution with sodium hypochlorite. 10) Add 1 volume of absolute ethanol, and vortex the mix. The volume is the same as the volume used to add AP3 solution. 11) Put the solution on the DNeasy minispin column, and centrifuge for 1 min at 10000 × g (to fix the DNA on the affinity column). The maximum volume of the column is 650 μl, so this has to be repeated several times. You have to discard the solution at the bottom of the Eppendorf tube after each step of centrifugation to avoid the overflow. 12) Place the column on a new 2-ml Eppendorf tube. 13) Pipette 500 μl of AW buffer on the column, and centrifuge for 1 min at 10000 × g (to desalt and wash the DNA). 14) Again, pipette 500 μl of AW on the column, and centrifuge for 2 min at 10000 × g (to be sure to eliminate all the buffer). Remove the AE buffer from the heater 2 min before use. 15) Place the column on a new 2-ml Eppendorf tube. 16) Pipette 100 μl of AE buffer on the column, and centrifuge for 1 min at 10000 × g (to elute the DNA). 17) Repeat this operation once. 18) Put the Eppendorf tube containing the DNA solution on ice (the column can be thrown away).
Polymerase Chain Reaction—
The PCR kit used is the PCR core kit from Qiagen. We have tested different Taq polymerase, and all are efficient on these primers. This choice is because this product is inexpensive and because of the presence of a premix of the four dNTP in one solution.
To make sure that the PCR will work for all of the teams, it is better to aliquot all of the solutions. This way you avoid contamination and false positives or negatives. To have enough material, put 15% volume in excess in each tube.
Each pair of students will amplify their purified DNA in the presence of the four different primers couples (four different PCR Eppendorf tubes). Tube A contains a mixture of chloroplast DNA-specific primers, tube B contains a mixture of lectin-specific primers, tube C contains a mixture of NOS terminator-specific primers, and tube D contains a mixture of 35 S promoter-specific primers.
To prepare these tubes, students label four PCR Eppendorf tubes. To each tube add, in order, the following: 33.9 μl of milliQ water (up to 50 μl), 5 μl of 10 × PCR buffer (containing 100 m M Tris-HCl buffer, pH 8, 500 m M KCl, 15 m M MgCl2), and 5 μl of the DNA solution prepared before. Mix gently, and place on ice.
To each tube, add 5 μl of the appropriate primer mix solution (initial concentration of 0.1 μg/μl). Mix gently and then return to ice.
To each tube add 0.6 μl of dNTP (a mixture of the four dNTP at 10 m M ) and 0.5 μl (1 unit) of Taq polymerase (QIAgen Taq polymerase core kit Qiagen). Mix gently and then return to ice. Keep the tubes on ice until the rest of the group has finished preparing their tubes.
Start the thermal cycler and verify the program. This program involves the following cycles. Step 1: 94 °C, 10 min for DNA denaturation Step 2: 94 °C, 1 min 63 °C, 1 min (annealing) 72 °C, 1 min (elongation) Step 3: 72 °C, 10 min for final elongation 4 °C (no time limit) for conservation of the PCR amplification products. Place the tubes in the thermal cycler, and start the thermal cycles as indicated. Step 2 is repeated for 34 cycles.
During the PCR, prepare a 3% w/v agarose gel in 0.5 × TBE. For this purpose, the students dilute 3 g of agarose in 100 ml of 0.5 × TBE electrophoresis buffer (0.09 M Tris, 0.09 M Borate, 1 m M EDTA), and boil the solution gently until it becomes clear. Pay attention to the overflow during boiling, which can alter the concentration. Then allow it to cool to ∼55 °C, and pour a gel.
Once the gel has solidified, the gel plate is placed on the electrophoresis apparatus and covered with 0.5 × TBE (to keep the same conductivity everywhere). This precaution avoids the dilution of the samples because of convection movements created by the buffer covering the gel after loading. At the end of the PCR, remove the tubes from the thermal cycler, and place them on ice.
In four fresh Eppendorf tubes, mix 8 μl of each amplified solution and 1 μl of 6 × loading buffer (0.3 M EDTA containing 10% glycerol to increase density, 0.25% bromphenol blue, 0.25% xylene cyanol, and 0.25% Orange G as migration indicator). In a fifth, mix 3 μl of 100-bp DNA ladder and 0.5 μl of 6 × loading buffer.
Onto the agarose gel, load 3 μl of 100-bp DNA ladder Promega solution with dye (which enables them to visualize fragments from 100 to 1500 bp), 8 μl of the chloroplast-amplified DNA solution with dye and then 8 μl of the lectin-amplified mix, 8 μl of the NOS terminator-amplified mix, and 8 μl of the 35 S promoter-amplified mix. Run the gel at 200 V until the Orange G (which migrates at approximately 50 bp) is at the bottom of the gel (∼30 min).
Stain the gel in ethidium bromide (to prepare the stain, put five drops of stock solution (10 mg/ml) in 200 ml of distilled water). (Be careful in handling ethidium bromide material safety data sheet number 1239–45-8, and use gloves and decontaminate the area used for this every evening.) Using a plastic support, transfer the gel into a box containing the solution of ethidium bromide. Stain for 10 min, destain in water for 1 min, and transfer the gel onto the ultraviolet transilluminator to visualize the bands of purified DNA. Take one photograph for each pair of students (Fig. 1). (Take care of exposure of the skin and particularly your eyes to the high intensity UV light. Use a Plexiglas screen.)
Paleofeces are valuable to archeologists and evolutionary biologists for their potential to yield health, dietary, and host information. As a rich source of preserved biomolecules from host-associated microorganisms, they can also provide insights into the recent evolution and changing ecology of the gut microbiome. However, there is currently no standard method for DNA extraction from paleofeces, which combine the dual challenges of complex biological composition and degraded DNA. Due to the scarcity and relatively poor preservation of paleofeces when compared with other archeological remains, it is important to use efficient methods that maximize ancient DNA (aDNA) recovery while also minimizing downstream taxonomic biases.
In this study, we use shotgun metagenomics to systematically compare the performance of five DNA extraction methods on a set of well-preserved human and dog paleofeces from Mexico (
Our results show that all tested DNA extraction methods yield a consistent microbial taxonomic profile, but that methods optimized for ancient samples recover significantly more DNA.
These results show promise for future studies that seek to explore the evolution of the human gut microbiome by comparing aDNA data with those generated in modern studies.
Lysis step during DNA purification - Biology
Priyanka Kardam 1 , Monica Mehendiratta 2 , Shweta Rehani 1 , Rashi Sharma 1 , Khushboo Sahay 1
1 Department of Oral Pathology and Microbiology, Sudha Rustagi College of Dental Sciences and Research, Faridabad, Haryana, India
2 Department of Oral Pathology and Microbiology, ITS Dental College, Greater Noida, Uttar Pradesh, India
Click here for correspondence address and email
|Date of Submission||30-Jan-2016|
|Date of Decision||17-Dec-2016|
|Date of Acceptance||16-Nov-2017|
|Date of Web Publication||18-Nov-2019|
Background: DNA analysis has a key role in forensic dentistry. However, techniques of DNA extraction and analysis are far from the reach of majority of medical professionals owing to its expensive set up. Aim: The present study was aimed at formulating a crude method of extracting DNA from human buccal mucosa cells using materials commonly available in the laboratory so that the medical professionals could get more exposure to molecular biology techniques. The objectives were to identify the DNA and to assess its purity. Methods: Buccal mucosa cells from 10 healthy volunteers were taken for DNA extraction following the protocol of cell lysis, purification, and precipitation. DNA was identified using standardized techniques like Diphenylamine test and its purity was assessed using a spectrophotometer. A gel electrophoresis apparatus was also constructed using readily available materials. Results: DNA was extracted from human buccal mucosa cells using a crude method. The standardized tests confirmed the presence of DNA contaminated with proteins. The locally made Gel electrophoresis model exhibited a faint halo around the wells instead of DNA bands. Conclusion: DNA extraction from human buccal mucosa cells was made possible using locally available materials and a crude method, but it was not of high purity.
Keywords: Buccal mucosa, DNA extraction, gel electrophoresis, molecular biology, spectrophotometer
|How to cite this article:|
Kardam P, Mehendiratta M, Rehani S, Sharma R, Sahay K. A crude method of DNA extraction and identification from exfoliated human buccal mucosa cells. Indian J Dent Res 201930:595-9
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Kardam P, Mehendiratta M, Rehani S, Sharma R, Sahay K. A crude method of DNA extraction and identification from exfoliated human buccal mucosa cells. Indian J Dent Res [serial online] 2019 [cited 2021 Jun 24]30:595-9. Available from: https://www.ijdr.in/text.asp?2019/30/4/595/271071
Molecular biology techniques are inaccessible to most medical professionals as these tests are expensive and require specialized equipment available only in some laboratories. These tests also require specially trained scientists and strict quality control. There is a general lack of practical knowledge about the modus operandi. 
The process of isolating nucleic acid consists of three steps, namely, cell lysis, purification, precipitation, and concentration. For cell lysis, the cell/nuclear membrane is disrupted to release the DNA. Purification involves removal and denaturation of proteins and inactivation of enzymes capable of degrading DNA. For precipitation and concentration, DNA and water bonds are broken and structural changes are induced to help DNA aggregate. The solution is then centrifuged at high speed to form a pellet. ,
The study was aimed at extracting DNA from human buccal mucosa cells using commonly available laboratory materials followed by its identification and assessment of its purity using standardized techniques such as diphenylamine (DPA) test and spectrophotometry. The objectives of the study were to extract DNA using different commonly available cell lysates (such as hand wash and liquid detergent) and purification agents (such as pineapple juice, contact lens solution and heat). Construction of a crude gel electrophoresis apparatus using readily available materials was also attempted.
This prospective study was performed in the Department of Oral Pathology and Microbiology, Sudha Rustagi College of Dental Sciences and Research, Faridabad. The sample group consisted of 10 healthy volunteers in the age group of 20 years who were chosen as the source of DNA. An informed consent of the patients and institutional ethical committee clearance were taken before proceeding with the study. The study was carried out in different phases namely DNA extraction, DNA identification, assessment of purity of DNA, and gel electrophoresis.
The first step was to collect the DNA extraction source, i.e. lightly adherent superficial epithelial cells or naturally exfoliated cells of oral cavity which were mainly from buccal mucosa. For the collection, a buffer for swishing the oral cavity was prepared by adding 1.5 g table salt (sodium chloride commercially available TATA salt) to 5.0 g baking soda (sodium bicarbonate commercially available TOPS India baking soda) and diluting it to a final volume of 120 ml with water. The volunteers were asked to take 10 ml of buffer solution and swish the oral cavity for 1 min. They were instructed to intentionally bite on the buccal mucosa so that more cells could be exfoliated into the buffer. The swished out buffer was then collected in a test tube.
The next step was to extract DNA from the collected epithelial cells present in buffer following the standard protocol of cell lysis, purification and precipitation. , For this study, we specifically used commercially available liquid hand wash (Lifebuoy total 10) and liquid detergent (Vim dishwash liquid). Using a pipette, 2 ml of cell lysate was added to swish out buffer in the test tube. The test tube was gently inverted 10 times, taking care to avoid foam formation. A similar procedure was performed using different varieties of cell lysates.
Ethanol (100%) was cooled (°C) by keeping overnight in the freezer, and it was used as the precipitating solution. Cooled alcohol of double the volume of solution in the test tube was carefully poured over it. The test tube was kept at an angle of 45° using a custom made stand in the entire process to ensure layer by layer deposition and no disturbance in the solution underneath. The test tube was again left undisturbed for 10 min at room temperature which was maintained at 20°C°C.
DPA test was used as a test for confirming the presence of deoxyribose (sugar moiety of DNA) in the spooled out strands from the sample. DPA stock solution was prepared by adding 15 g DPA to acids (1000 ml glacial acetic acid and 15 ml sulfuric acid). The working solution was freshly prepared before use by the addition of 5 ml of 2% acetaldehyde (100 μl acetaldehyde and 5 ml water) for each 1000 ml of the stock solution. Solution was kept in dark glass to avoid its darkening by light. To 1 ml of DNA sample, 2 ml of DPA working solution was added and the test tube was kept in boiling water for 15 min.
Assessment of purity of DNA
Spectrophotometer was used to measure the absorbance (wavelength at which the light is absorbed) which helped in the assessment of purity of DNA in the sample obtained. The spectrophotometer (HITACHI U-2900) was switched on for 10 min for the lamp to warm up followed by calibration of the instrument according to the standard protocol. DNA sample was taken in a quartz cuvette, and absorbance was measured at 260 and 280 nm for different combinations of cell lysates and purification enzymes. A260/A280 ratio was calculated to estimate the purity of the sample.
A running buffer was prepared by adding 0.05 g of table salt (Sodium chloride), and 2 g of baking soda (Sodium bicarbonate) to 1 L distilled water. The pH was brought to 7.5 using a pH meter. Next, the cooled glass plate with agar gel was placed into the plastic box (electrophoresis chamber) and the ends of copper wires tied to glass plate were then connected to a series of 9 volts batteries. Running buffer was poured into the plastic box until the agar gel was immersed up to 3 mm in it.
Loading buffer consisted of 0.5 ml glycerine mixed with 0.1 ml distilled water. This loading buffer can be used as to 吆 dilution for the DNA sample, i.e. for 0.5 ml of DNA sample, 0.05.1 ml of loading buffer was used. After this, DNA sample with loading buffer was poured into the well cut in the agar plate using a micropipette. In another well, orange G (negatively charged) dye was added as control. This setup was left undisturbed for 45 min.
To visualize the DNA bands, staining was done by pouring methylene blue solution (0.1%) over the agar gel. The setup was left undisturbed for the whole night.
DNA (Deoxyribonucleic acid) provides a unique identity to every human being. It is an integral component of all tissues of body such as hair, nails, biopsy sample, saliva, blood, and buccal mucosa. These sources show variations only in quantity and quality of the DNA.
There has been a great progress in the field of forensic science owing to the development of newer and more reliable techniques of DNA extraction and typing. ,, However, the molecular biology is still at the grass-root level owing to the expensive equipment and specialization it demands with regard to technology.  This study was conducted with an aim to bring the usually unreachable molecular biology. A pilot run based on MacGyver's experiments to extract DNA was carried out on different fruits and vegetables such as onion, kiwi, and banana following which the study was performed on human buccal mucosa cells.
On the cells of human buccal mucosa, the study was carried out in four stages, namely, DNA extraction, DNA identification, assessment of purity of DNA, and gel electrophoresis. A successful DNA extraction was carried out following the routine protocol of cell lysis, purification and precipitation using cheaper and easily available substitutes of expensive chemicals and equipment. The next step was to confirm the presence of DNA in the solution. For this purpose, the biochemical test for the presence of deoxyribose was used. DPA reagent confirmed the presence of DNA by reacting with deoxyribose (sugar moiety of DNA) in an acidic medium to produce a blue colored solution. 
To assess the purity of DNA extracted, spectrophotometry was used. Spectrophotometry is based on the principle of Beer-Lambert law, which states that when a sample is placed in the beam of a spectrophotometer, there is a direct linear relationship between the amount (concentration) of its constituent (s) and the amount of energy it absorbs. The wavelength at which the light is absorbed (absorbance) is a function of molecular structure of the compound.  The nitrogenous bases present in the DNA, allows it to absorb light in the UV range (260 nm). Proteins are the most common contaminants extracted along with DNA and they also absorb in the ultraviolet range (280 nm). To determine the purity of DNA sample extracted, a ratio of absorbance at A260 nm and A280 nm is used. A good quality DNA sample should have A260/A280 ratio of 1.7.0. If the A260/A280 ratio is more than 1.75, DNA is pure enough to proceed. If the ratio is ɮ.1, it is not DNA and if it is ə.75, the DNA is highly contaminated with protein. 
In the present study, an attempt was also made to construct a working model of gel electrophoresis apparatus. The movement of charged particles through an electrolyte when subjected to an electric field resulting in their migration towards the oppositely charged electrode is known as electrophoresis.  Electrophoresis is affected by various factors such as net charge on the particles, mass and shape of the particles, pH of medium, and strength of electrical field. Agarose gel electrophoresis is a routinely used method for separating proteins, DNA or RNA.  Gel electrophoresis separates DNA fragments by molecular weight in a solid support medium (agarose gel). DNA samples are poured into the sample wells using a pipette and application of an electric current at the anodal end (negative) causes the negatively-charged DNA to migrate (electrophorese) toward the cathodal end (positive). The rate of migration is proportional to size: smaller fragments move more quickly and move farther away from the anodal end. In our study, this model was not fully successful this could probably be due to the use of an inferior quality replacement of agarose. 
For better results the authors suggest the use of better/known substitutes available in the market, for example, agarose instead of agar, proteinase K instead of pineapple juice and voltage regulator instead of transistor batteries. However, more sampling sites could also be tried for DNA extraction, for example, blood, exfoliated cells from different sites of the oral cavity.
Hence, to conclude we would like to say that molecular biology is still at grass-root level in India which leads to lack of knowledge amongst the health-care professionals and students. However, if the researchers pour in the efforts to devise cost economic techniques for performing molecular biology, it could lead to an increase in exposure and thus awareness of these professionals.
The authors would like to acknowledge Late. Dr. P. K. Jain without whose guidance this project would not have been possible. We would also like to acknowledge Dr. T. K. Mishra whose help was very fruitful for the project.
In this study, when either a direct lysis or cell extraction method was used to isolate DNA from soil samples, humic acid-like substances were observed in all crude DNA extracts. Indeed, it was also observed that additional cell extraction steps increased the content of humic acid-like substances in the cell pellet (data not shown). Tsai & Olson (1992) showed that the presence of 27 μg humic acid-like substances or 10 ng pure humic acid is sufficient to inhibit subsequent polymerase chain reaction (PCR). Additionally, Steffan et al. (1988) showed that the presence of 10 μg humic acid would strongly interfere with the hybridization between probe and target DNA in dot blots, even when the latter was at 100 ng. In the present study, attempts to remove these humic acid-like substances by gel filtration failed. When subjected to caesium chloride–ethidium bromide (CsCl-EtBr) density gradient ultracentrifugation, however, DNA of suitable purity was obtained, as demonstrated by their restriction enzyme digestibility and A260/A280 ratios. This purification technique was therefore used in the subsequent parts of the study it does, however, have the disadvantage of being relatively time-consuming.
When the blending methods (methods 6 and 7) were used to extract bacteria from sandstone shale alluvial soil, about 50–80% of the culturable cells in the sample were isolated ( Table 1). In a similar study, Holben et al. (1988) reported recovery rates of about 35% whether based on viable count or direct count data. Jacobsen & Rasmussen (1992) failed to find any significant difference between the bacterial recovery percentages of cation-exchange resin (method 5 here) and the blending methods. In the present study, however, Chelex-100 recovered only about 30% of the total aerobic bacteria, i.e. somewhat less than either of the blending methods ( Table 1). Furthermore, although purity was high, the Jacobsen and Rasmussen method (method 5 in Table 1) yielded the smallest quantity of DNA. It has been reported that the amount of DNA obtained by the direct lysis method can be 20–70 times higher than by the cell extraction method ( Steffan et al. 1988 ). In the present study, the direct lysis method described by Holben (1994) produced yields from sandstone shale alluvial soil that were at least an order of magnitude better than from the three cell extraction methods ( Table 1). This method was then used to extract DNA from Taiwan clay and sandstone shale and slate mixed alluvial soil receiving different kinds of fertilizer. Higher amounts of DNA were extracted from soil receiving organic fertilizer than from the same soil receiving chemical fertilizer ( Table 2). This is consistent with reports that soils managed under an organic farming regime generally contain higher microbial biomass ( Fraser et al. 1988 Kirchner et al. 1993 ). When sandstone shale alluvial soil was used, both yields of DNA and cfu counts were generally quite low when compared to previous reports. It is believed that this reflects the soils used rather than the extraction protocols.
In the total DNA prepared from soil samples, fragments of various molecular sizes were found ( Fig. 1). When E. coli was added to acid-washed sea-sand and subjected to the same extraction procedure, however, no fragmented DNA was detected. This suggests that Holben’s direct lysis method itself caused minimal damage to the structural integrity of the extracted DNA, and that the small-size DNA fragments in the total DNA extracts could be interpreted as extracellular DNA at various stages of degradation. Evidence of degradation of the large-molecular-size DNA fraction was also seen when soil samples were stored at 4 °C for several weeks ( Fig. 1, Lanes 2–9). Currently, the reason for this is unclear. In order to minimize such DNA fragmentation, it is therefore recommended that the total DNA should be extracted from the soil sample as soon as possible after collection.
When a direct lysis method is used to extract bacterial DNA from soil samples, it is possible that the DNA of eukaryotic organisms (e.g. fungi) might also be extracted. Steffan et al. (1988) , for example, were unable to confirm the absence of eukaryotic DNA in their DNA extracts. In the present study, a weak hybridization signal was observed between probes made from total soil DNA and the genomic DNA of S. cerevisiae but not with the other eukaryotic target DNA tested. This suggests that the DNA extraction method used (i.e. Holben’s direct lysis method) did not, in fact, extract fungal DNA. Further evidence was provided by subjecting fungal hyphae to Holben’s direct lysis, with or without the addition of sandstone shale alluvial soil no nucleic acid was detected in extracts obtained from samples containing F. solani, while RNA, but not DNA, was detected in extracts obtained from samples containing P. aphanidermatum ( Fig. 2). The presence of RNA was further confirmed by treating the samples with RNase. This difference can be explained by the fact that Fusarium hyphae are septate while Pythium are not, so that any damage to the hyphae of P. aphanidermatum would result in a massive release of cytoplasm. This result is interpreted as indicating that while the direct lysis method described by Holben can cause some damage to the fungal cell wall, it is nonetheless not severe enough to cause subsequent release and lysis of the fungal nucleus.
It is concluded that the direct lysis method described by Holben (1994) , together with CsCl-EtBr density gradient ultracentrifugation, can yield the greatest amount of high quality DNA.