DNA extraction of Molluscs (long protocol)

(Maureen Krause’s protocol modified for eppendorf tubes by Elizabeth Boulding’s lab).

This protocol works for most species of molluscs (Dreissena, Littorina) we have tried it on.  We don’t use it as often as we used to as it takes nearly 3 days to do all the steps.  Instead we usually use our modification of the BioRad Chelex protocol or one of the 96 well plate versions of the commercial DNA extraction kits.

We have found that careful preparation of DNA templates is essential if people are to enjoy their laboratory work with these slimy invertebrates.

I. Tissue Preparation:

When extracting DNA from gastropods visceral mass is used, when extracting from veliger larvae the entire animal is used, and when extracting from bivalves the adductor muscle is used.  The samples are brought out from their storage in the –80o freezer and they are allowed to thaw on ice.  If using ethanol preserved tissue, the tissue is soaked for several hours in water in the refrigerator.  Ideally the water is changed every hour or so, and then the molluscs are blot-dried on a paper towel.

A large glass plate divided into 18 separate squares is washed with 10% HCl and it is then rinsed with distilled water and allowed to air dry.  The glass plate may be found by the filing cabinet at the back of the lab.  Before proceeding, the snail hammer and two pairs of forceps are brought out and placed on a paper towel near the glass plate.  Also three small 100 ml beakers are brought out.  One is filled with approximately 50 ml of 10% HCl, and the other two are each filled with approximately 50 ml of distilled water.  Next, one snail is placed into square #1 on the glass plate and the hammer is used to gently crack its shell.  For thin shelled gastropods one light tap is generally enough to do the trick.  Then the forceps are used to pry the shell from the soft tissue of the snail, and when working with small snails it is useful to rip and crush the soft tissue a bit in order to expose enough DNA.  Beware not to damage the tissue too much, otherwise DNAses will be released and they will degrade the DNA, however, keep in mind that you want to separate all the soft tissue from the shell.    To clean the forceps between animals:

1) The tools are rinsed in the distilled water found in the 100 ml beaker

2) The tools are washed in 10% HCl solution, found in the 100 ml beaker, for 30 s

4) The tools are rinsed well in the distilled water found in the second 100 ml beaker

5) The tools are blot-dried on a paper towel

Be sure to dry the instruments well before using them on the next specimen, otherwise you may find that the acid will corrode the instruments and contaminate the tissue samples.  Proceed to place the next mollusc on square #2 on the glass plate once you are done with the first one.  Separating the molluscs into the squares is important because it helps to prevent DNA contamination.

II. Lysis and Digestion with Proteinase K:

No more than 5-10 mg of wet tissue (weigh it or you’ll use too much!) is added to a 1.5 ml microcentrifuge tube containing a “digestion” solution of:

336 ml ultrapure ddH2O (Front sink)

20 ml 20X SSC (Large 4 oC fridge)

44 ml SDS (20% detergent solution) (Shelf - liquid chemicals)

0.2 mg Proteinase K (Large 4 oC fridge in a desiccated  tupperware)

Note:  This recipe will make enough solution for one tissue sample.  Multiply by the number of tissue samples you will be extracting in order to get the correct quantities. 

The 20X SSC is usually found in the large 4oC fridge in the back room.  If there is none there, you may want to check the small fridge by the fume hood, but if there is none there either, you will need to make some.  In order to make 20X SSC, dissolve

175.3 g of 0.15 M NaCl (Shelf - solid chemicals) and

88.2 g of 0.015 M Trisodium citrate (make sure it’s really trisodium!) (Shelf - solid chemicals) in

800 ml of ddH2O (Front sink)

Adjust the pH to 7.0 with a few drops of a 10N solution of NaOH.  Adjust the volume to 1 liter.  Then dispense into aliquots and sterilize by autoclaving.

We usually make up enough “digestion” solution for 18 tubes (one centrifuge full) at once and leftover solution may be frozen.  The samples are incubated for up to 16-20 hours at 55oC in the water bath by the front of the lab room.  You may gently invert the tubes occasionally to speed things up.  When the digestion is complete the solution should be clear and there should be no lumps except for pieces of shell.  If there are lumps then something may be wrong with your chemicals. 

III. Removal of Proteins with MCIA as the organic solvent:

The proteins are removed using a methylene chloride/isoamyl alcohol mixture (MCIA is by volume 24:1 methylene chloride:isoamyl alcohol) as the organic solvent for the extraction.  MCIA is cheaper and better than phenol/chloroform for most molluscs but it is also more toxic.  Keep the mixture in a good fume hood and on ice to minimize evaporation and use gloves, a lab coat, and safety glasses when working with the chemical.  Reading the chemical safety data sheets will encourage you to be careful.

1.     130 ml of 5M NaCl (Shelf - solid chemicals) is added to every sample and the tube lids are shut in order to invert the tubes 5 times. 

2.     44 ml of MCIA (Small 4oC fridge) is added to every sample and the tubes are inverted 5 times.  The lids are opened to release the pressure and then they are closed. 

3.     The tubes are left on ice for one to two hours at the end of which the lower layer has formed into a “gel”. 

4.     The samples are centrifuged (preferably at 4oC) at 2000g (check the microcentrifuge manual to convert rpms to g) for five minutes and then at 6000g for ten minutes. 

5.     The supernatant (= upper layer) is removed carefully using a 200 ml pipette. You should always use wide bore plastic pipette tips or cut 1.0 mm off the end of the standard tips to avoid shearing the genomic DNA.   Don’t be greedy!  For really clean DNA make sure none of the intermediate, mixed layer is taken up into the pipette tip.  The supernatant (= upper layer) is transferred to a new set of test tubes.  It is useful to label the new test tubes with a permanent waterproof marker and to line up the new test tubes beside the older ones in order to avoid placing supernatant in the wrong tube.

6.     The supernatant is placed in another tube containing 440 ml of MCIA.

7.     The tubes are inverted 5 times and then the lids are opened to release the pressure before they are closed for the next step. 

8.     The samples are centrifuged at 6000g for five minutes. 

If you would like to obtain really clean DNA you may repeat steps 4-7 (organic extraction) again.

1V. Ethanol Precipitation of DNA:

1.     The supernatant is removed and placed in a tube containing two volumes of 95% cold ethanol (-20oC fridge).  The ethanol we have in this lab is not quite 95% even though the label indicates that it is.  It is closer to 90%.  Therefore when adding the cold ethanol, do not add only two times the volume, add instead 2.2 times the volume of the supernatant. 

2.     The tubes are inverted 10 times before being placed in the -20 oC  fridge to precipitate for a minimum of three hours.

3.      The solution is spun down at 10,000g (max speed) for 30 minutes.  It is useful to place the tubes in the centrifuge in such a way that the lids of the tubes are all facing towards the center of the centrifuge.  This will make spotting the DNA pellet easier.

4.     The ethanol is carefully removed with a pipette so that the DNA pellet is not disturbed.  At times the DNA pellet is very small and difficult to see.  Often it may be found attached to the side of the tube near the wall that is facing outward when you put the tube in the centrifuge.  Other times the pellet will be free-floating in the alcohol, and this makes extracting the alcohol without disturbing the pellet a little more difficult.  You may wish to use the magnifying glass with light attached in order to make sure that you do not accidentally lose your pellet.  Normally, a second set of empty tubes is set up for the disposal of the ethanol.  Therefore if you should accidentally suck up your pellet with the ethanol, you may retrieve it from the “waste” test tube.

5.     The DNA pellets are washed by adding 500 ml of 70% cold ethanol (-20oC fridge) to the tubes. 

6.     The tubes are centrifuged at 10,000g (max speed) for 5 minutes.

7.     Again, the ethanol is carefully removed with a pipette so that the DNA pellet is not disturbed.

8.     The pellet is air dried until just barely dry but with all traces of ethanol gone.  Normally the tubes are allowed to dry on their sides face down on a slanted eppendorf tube holder wrapped in Kim Wipe.  Do not let the samples dry overnight because the pellet may cake on to the wall of the tube and never resuspend in water.

If you do not see any DNA pellet at step 4, then you may wish to add some NaCl to the sample and store it in the –20oC fridge for at least three hours in order to encourage precipitation.  You may then start the ethanol precipitation again.   

V. Resuspension and Dilution of DNA:

Resuspend the pellet in 50 ml sterile (PCR quality) distilled water and hold the tube up to a bright light to confirm the pellet has really resuspended.  Heating the tube in a waterbath at 50 oC for up to 30 minutes (DO NOT heat above 55 oC) is helpful for stubborn DNA that refuses to resuspend. The template solution is diluted 1:50 for routine applications of the polymerase chain reaction. The ideal dilution is 20 nanograms per microlitre for sensitive PCR applications such as microsatellites.  Even if DNA concentration is low, the dilution must be at least 1ul of the DNA concentrate in 50 ml distilled water so that the polysaccharide contaminants are low enough in concentration to avoid inhibition of PCR.  The diluted DNA template may be stored at 4 oC for a month or so.   However, the remaining concentrated DNA concentrate should be stored at -70oC so it can be thawed on ice and more dilutions made when the original diluted DNA solution stops working in PCR reactions.