Fast and Sustainable Thermo-osmotic DNA Extraction Protocol for Trans-spectrum Contingency and Field Use

Materials and reagents

The provisions below do not include expendables and instrumentation necessary for PCR and downstream procedures, such as gel electrophoresis.

Biological materials

1. Swab/scrap samples from human oral mucosa (tongue or inner cheek/gum)

2. Fungal mycelia grown on fruits and vegetables

Reagents

1. Commercially available distilled water for house use (i.e., steam ironing) from any convenience store and manufacturer, upon availability

2. Absolute ethanol, 99.8% denatured with IPA, MEK, and Bitrex pure (Panreac/Applichem, catalog number: 147194) or any other of comparable concentration (high purity is not essential)

Solutions

1. 70% ethanol (see Recipes)

Recipes

1. 70% ethanol

   Reagent	Final concentration	Volume
   Ethanol (absolute)	70%	700 mL
   H2O	30%	300 mL
   Total	n/a	1,000 mL

Laboratory supplies

1. Falcon-shaped plastic tube, 50 mL (KIMA Vacutest, catalog number: KIMA-17102-50)

2. Beaker, 1,000 mL (Hamed, catalog number: 303173)

3. Polypropylene floating rack, 8 positions (Flinn Scientific, catalog number: FB1671)

4. Pasteur pipettes, glass (Hamed, catalog number: 303107)

5. Rubber bulb for Pasteur pipettes (Isolab Laborgerate, catalog number: 084-03-001)

6. Pipette P20, 0–20 μL (Gilson PIPETMAN^®, catalog number: F123600)

7. Pipette P200, 20–200 μL (Gilson PIPETMAN^®, catalog number: F123601)

8. Pipette P1000, 200–1,000 mL (Gilson PIPETMAN^®, catalog number: F123602)

9. Pipette tips, 0–200 μL (KIMA Vacutest, catalog number: KIMA-18260)

10. Pipette tips, 1,000 μL (KIMA Vacutest, catalog number: KIMA-18172)

11. Plastic microtubes, 1.5 mL (Pierce^TM, catalog number: 69715)

12. Plastic microtubes for PCR, 0.2 mL (Thermo Scientific^TM, catalog number: AB0620)

13. Scalpel blades (no. 22 stainless disposable; FEATHER, lot 03067430) or box cutter (e.g., Q-Connect, catalog number: 233471)

14. Rack (50-position for 18 mm diameter microtubes) (Fruugo)

15. Hydrophilic cotton (e.g., pharmacy package)

16. Plastic spoons (e.g., any convenience store, 10 pc package), spoons (e.g., stainless steel 18/0 convenience store), glass slides (76 mm × 26 mm, Knittel Glass, catalog number: 303157), or wood tongue depressor (disposable, Heine, catalog number: 508424)

17. Plastic pellet pestles, blue polypropylene autoclavable (Sigma-Aldrich, catalog number: Z359947-100EA)

18. Cotton (any pharmacy/convenience store) or paper tissue/napkins (any convenience store)

Equipment

1. Camping gas stove (e.g., KEMPER, 12.3 cm × 12.3 cm × 21.7 cm, model: 06450208) or electric kitchen stove burner (e.g., Severin, model: 3519368)

2. Lighter (e.g., KEMPER, model: 10422)

Procedure

1. Sample collection (see General note 1) and preparation

   1. If swab/tissue collector item is not sterile, rub it with 70% alcohol on a piece of cotton or paper tissue/napkin and light it with the lighter.

   2. Allow the flame to die out and let it cool down in the air.

   3. Sample gathering; either:

      1. Collect a swab from the upper surface of the tongue or the inner surface of the cheek by mildly scrapping with any of the proposed or other similar objects (see General note 2);

      2. Cut off a piece (3–5 mm in diameter) of the mycelium grown on the plant tissue. Alternatively, scrap it from animal tissue or scrap bacterial or yeast growth from any such surface by using disposable sterile scalpel bladed or flame-sterilized scalpel, razor, or box cutter (see General note 3).

   4. Store sample in a 1.5 mL microtube (see Troubleshooting 1) until step B1. If needed, use some guidance/assistance to put it in (see General note 4).

   5. Fill a beaker to half capacity with tap water; this will serve as a water bath.

   6. Fill a 50 mL Falcon tube to half capacity with commercially available distilled water and place it into the beaker.

   7. Heat the beaker on the stove (camping gas or electrical) at full intensity until the water bath starts boiling; then, reduce to half intensity to keep the water boiling during the entire procedure.




2. Cell lysis

   1. Add boiling hot water to the extraction tube

   1. If mechanical assistance in lysis is required (due to the nature, quantity, or physical qualities of the sample), add 250 μL of boiling-hot distilled water or half the volume you intend to use (precision is of no particular importance) to the extraction microtube containing the sample from step A4 (see General note 5).

   2. If no mechanical assistance is required, add the sum of the extraction volume of boiling water and go straight to step B4. Caution: Steam may come out of the Falcon tube upon unscrewing and the tube may be hot. Take precautions during touching and handling and keep it safely placed on the respective rack or any similar support (e.g., a small beaker or glass).

   2. Crush with a spiral motion of the pestle or other similar instrument for 2–3 min.

   3. Use an equal volume of boiling-hot distilled water (e.g., 250 μL, as suggested in step B1a) to wash cellular debris of the pestle within the microtube.

   4. Close the lid of the microtube and vortex or shake vigorously by hand for 1 min (see General note 6)




3. Incubation

   1. Place the microtubes in the boiling water bath for 20–30 min or more, according to the extraction volume and rigor of extraction (see General note 7; see Troubleshooting 2). Use the incubation time to prepare the molecular testing reactions, if applicable (step E1).

   2. Upon completion, carefully open microtubes and pipette 10 μL from the center of the extraction volume to a PCR microtube.

      Caution: Steam pressure will have built up in the microtube; thus, handle with care and away from the face. Preferably, place in a rack, either custom made or improvised, before opening and further handling.

   
   

4. Dilutions

   1. Transfer 5 μL of the 10 μL to the first dilution tube and mix with 45 μL of distilled water by softly pipetting in and out 5–10 times, so as to have a 1/10 dilution. Use a 20 μL pipette for the whole procedure.

   2. Remove 5 μL of the 1/10 dilution to a second PCR tube (of 200 μL volume or as used per standard practice).

   3. Remove another 5 μL of the 1/10 dilution to the second dilution tube and mix with 45 μL of distilled water by softly pipetting in and out 5–10 times, so as to have a 1/100 dilution.

   4. Remove 5 μL of the 1/100 dilution to a third PCR tube (see General note 8).

   
   

5. PCR amplification

1. Prepare the mastermix, according to the desired protocol, while incubating the extract in the water bath and keep on ice until use (see General note 8). This step should be implemented during step C1.

2. Dispense the required volumes of mastermix to the three PCR tubes with the serial dilutions for each sample and proceed to amplification in thermocycler as per standard procedure/protocol. This step should be implemented after step D4.

Data analysis

Each extraction was performed in triplicates independently, with no more than a week between two successive efforts. Any of the three dilutions producing amplicon was deemed successful. Here, we report that the protocol can be used in cases where at least two of the three independent extraction efforts produced amplicon in at least one out of three serial dilutions (the undiluted crude extract being understood as the first in the series of the three dilutions).

Validation of protocol

There is no way to verify the performance of the present protocol directly: it is an extraction protocol and since its product is not purified, it cannot be tested properly—neither by spectrophotometry, as the cell debris would absorb or scatter the emitted UV, nor by gel electrophoresis, as said debris would obstruct the motion of the DNA, mar the electric field, and also absorb the fluorescent dyes, creating noise and smear. The only applicable validation approach is indirect, by the success or failure of some downstream nucleic acid amplification test (NAAT). To secure validation, negative controls with distilled water-only added in equal volume to the template were used, to ensure that amplicon production was not due to contamination of the NAAT reagents, expendables, or equipment.

Furthermore, to ensure that the NAAT (here, PCR) does work in terms of protocol, cycler, and reagents, positive controls were used. These were semi-simultaneous extractions of mycelial tissue, by other established methods such as alkaline extraction and/or CTAB (Velegraki et al., 1999a; Goudoudaki et al., 2021), and also of stock DNA extracts kept in the lab from previous projects, extracted with conventional protocols and occasionally quantified by mini-spectrophotometer (Psarias et al., 2020). The full procedure for validation and controls may be found in the source publication (Goudoudaki et al., 2023). In short, from a number of tried applications, positive PCR results were returned for two cases of samples: (a) swab samples tested either by consensus sequence PCR targeted at the local bacteriome (product band of size compatible with the expected target range) or for one specific endogenous (human) locus (product band of the expected size); (b) consensus sequence PCR on extracts of fungal mycelia grown on the surface of fruits held in refrigeration (product band of size compatible with the expected target range).

General notes and troubleshooting

General notes

1. The method is valid with relatively soft and uncomplicated samples in which the target genome is abundant (highly enriched). Mouth swabs produced specific human genome amplicons and consensus sequences amplification product for bacteria but not for fungi, which are much less populous in the oral mucosa.

2. Glass slides for microscopy, plastic or metallic spoon—especially disposable ones for deserts—or the handle of all-metallic cutlery or tongue depressors may be used.

3. During sampling, samples as pure as possible of target cells must be collected (pure is understood as “containing as few admixtures and as homogenous a cellular population as possible”). The quantity of the sample must be limited, as there is no purification step to dispose of cell debris and consequent background noise and possible inhibitions.

4. If needed, help/guide the sample within the microtube with a similarly sterilized microbiology needle or a similar piece (i.e., a second glass slide) or a plastic tip.

5. If mechanical assistance is required, take caution that the volume of sample + boiled water is approximately 500 μL. Additional boiling water will be added to wash off the pestle from attached debris, but it is better that the final volume does not exceed 1 mL.

6. If the extraction microtube is too full (> 1 mL), punch holes using a needle, thin nail, or pin in the microtube lid before incubation so the pressure will not blow the lid open, throwing the content out of the tube.

7. Use purpose-built floating rack or punch/cut holes to felizol (EPC–Expanded PolyStyrene) sheets of less than 50 mm thickness recovered from packaging boxes.

8. Functional concentration during the NAAT may be adjusted to other values: either by i) increasing the template volume in the reaction tube while subtracting an equal volume of sterile distilled H₂O from the mastermix or ii) if no mastermix is prepared, by changing the volumes in the reaction microtube. Occasionally, the opposite may also be applicable (diminishing the template volume while increasing that of the sterile distilled H₂O). Similar manipulations are applicable at the serial dilutions step.

Troubleshooting

1. If the sample size within the microtube is excessively large (operative’s call, depending on the difficulty to properly handle the extraction mix, but see General note 6 for an example), it may be divided in two. If it is solid, this can be done by cutting with a sterilized surgical disposable blade; if it is liquid, the excess may be pipetted off to another tube, creating a technical replicate.

2. Longer incubation time is advised (a) when treating larger samples in terms of volume/mass and (b) if the cell lysis phase is less rigorous, as in the absence of mechanical assistance. Refer to step C1.

Acknowledgments

This project was in part funded by the European Commission, Horizon 2020 (H2020-668353; Ubiquitous Pharmacogenomics).

This protocol is based on the research paper (Goudoudaki et al. 2023).

Competing interests

The authors declare no conflicts of interest or competing interests.

Ethical considerations

The study was approved by the Institutional Research Ethics Committee of the University of Patras. Swabs were collected from the operatives performing the development of the protocol and the subsequent analysis as explicitly stated in the paper (Goudoudaki et al., 2023).

References

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