16. Protocols used in Cuming Laboratory

The following protocols are established by Dr. Andrew Cuming and

Dr. Yasuko Kamisugi in the University of Leeds.
Stock solutions for moss media

Solution B

MgS0₄.7H₂0 25 g (anhydrous MgS04 12 g)

distilled H₂0 to 1 l

Solution C

KH₂PO₄ 25 g

distilled H₂0 500 ml

Adjust pH to 6.5 with minimal volume of 4 M KOH

Make up to 1 l with additional distilled H20

Solution D

KNO₃ 101 g

FeS0₄.7H₂0 1.25 g

distilled H₂0 to 1 l

Over time, this solution develops a red precipitate. Shake thoroughly to resuspend this immediately before removing an aliquot.

Trace element solution (“TES”)

H₃BO₃ 614 mg 　　　　　 MnCl₂.4H₂0 389 mg　　　　　　　　 AlK(S0₄)₂.12H₂O 110 mg

CoCl₂.6H₂0 55 mg　　　 CuSO₄.5H₂0 55 mg 　　　　　　　 ZnS04.7H20 55 mg

KBr 28 mg 　　　　　　　　 KI 28 mg　　　　　　　　　　　　　　　　 LiCl 28 mg

SnCl₂.2H₂0 28 mg　　　 Na₂MoO₄.2H2O 25 mg 　　　　　　　　　　　 NiCl₂.6H₂0 59 mg

distilled H20 to 1 l

We use this trace element solution but others have used other recipes, the exact composition of the trace element solution is probably not important.

0.5M diammonium tartrate 　 9.2 g made up to 100 ml with distilled H₂O

These solutions can be stored at 4 C without autoclaving.

IM CaCl₂ (autoclaved) 　　　　　

Store in 10 ml aliquots

For routine culture of moss tissue, we use “BCD” medium containing 5mM diammonium tartrate and 1mM CaCl₂, solidified with agar.

Solution B　　　　　　　　　　　　　　　　　　　　　 10 ml

Solution C　　　　　　　　　　　　　　　　　　　　　 10 ml

Solution D　　　　　　　　　　　　　　　　　　　　　 10 ml

0.5M diammonium tartrate　　 10 ml

Agar (Duchefa plant agar)　　　 5.5 g

Distilled H₂O　　　　　　　　　　　　　 to 999ml

Autoclave, and add 1ml sterile 1M CaCl₂ immediately before pouring into petri plates.

We use Sarstedt 9cm diameter unvented petri plates. The use of unvented plates reduces evaporation and drying out of cultures.

If diammonium tartrate is omitted, a greater proportion of caulonemal filaments develop.

Diammonium tartrate should be omitted for culture of plants when sporophyte development is desired.

Other supplements that can be used include:

Growth Supplements

substance 　　　　　　　　　　　　　　 concentration 　　　　　　 weight 　　　　　 stock

　　　　　 in the medium 　　　　　　　　　　　 per litre 　　　　 sol’n (ml)

p-aminobenzoic acid 1.8 µM 　　　　　　　　　　　　　　　　　 247 µg 　　　　　 1000

D-glucose 　　　　　　　　　　　　　　　 0.5% (w/v) (= 28 mM) 5 g 　　　　　　　　 100

nicotinic acid 　　　　　　 8 µM 　　　　　　　　　　　　　　　　　　　 1 mg 　　　　　　　 1000

D-sucrose 　　　　　　　　　　　　　　 0.5% (w/v) (= 15 mM) 5 g　　　　　　　　　 100

thiamine HCl 　　　　　　 1.5 µM　　　　　　　　　　　　 0.5 mg 　　　　　 1000

(Note: The vitamin supplements are only used for culture of auxotrophic mutants; and sugar supplements for growth in darkness. For routine culture these are not used.)

“BCD” plus NH₄medium is used for routine culture of moss as spot inocula and as homogenate cultures. It promotes rapid growth. In homogenate cultures, the tissue will be almost exclusively chloronemal filaments for the first week.Subsequently, caulonemal filaments will differentiate and buds will form, giving rise to gametophores.

BCD lacking NH₄ will favour caulonemal filaments, and spot inocula grown on this medium will eventually undergo sexual reproduction to generate spore capsules by self-fertilisation. This does NOT occur on medium containing ammonium ions.

Cellophanes

Cellophanes discs are Type 325P, obtainable from AA Packaging Ltd., Liverpool Road, Walmer Bridge, Preston, Lancashire PR4 5 HY, England. At one time you had to purchase a minimum order of 50,000 discs (!). This is no longer the case, but you still have to order a lot, and pay in advance (about ￡8 per 1,000 discs).

We sterilize cellophanes by interleaving them with 9cm filter discs (e.g. Whatman No 1) – usually about 30 discs at a time in a glass petri dish. (The filter papers can be reused many times!)

Once sterilized, the cellophane discs can be overlaid onto agar medium: pick up a cellophane disc using two pairs of forceps. Hold it in a “U”-shape and touch the bottom down onto the agar. Gently lay the disc onto the agar. – The disc may wrinkle at first. Don’t worry. Once it has thoroughly wetted　 any wrinkles and trapped air bubbles can be removed.

Plates can be overlaid with cellophane and stored at room temperature for several weeks (although it is generally advisable to prepare plates mor or less as you need them).

Homogenate plates

3

2

Homogenised moss is used for continuous vegetative propagation

6

5

4

1

9

10

8

7

　

13

12

11

　　　　　　

Moss utensils: sterile 10ml pipette, spatula, forceps, homogenizer unit.
Harvest moss by scraping from cellophane with flat end of spatula
The moss is drawn easily into a clump
Pipette 10ml sterile water into the Universal bottle
The moss is easily picked up with forceps…
…and dropped into the water
The blade unit is put in place.
Our ancient 1970s homogenizer
The homogenizing unit is clamped in place
Homogenise
Fragments generated by homogenization.
Rapidly dispense 2ml homogenate onto a fresh plate
This is what it looks like.

Plates are incubated at 25 C under continuous illumination. After a week, you should have a lawn of filamentous culture like that in panel 1. Always pipette a small aliquot of homogenate into a vial of L-broth that can be placed in the tissue culture room alongside the plates, to monitor contamination.

The nearest commercially available equivalent to our type of homogeniser that I have seen is the IKA Ultra-Turrax Tube Drive homogenizer:

(http://www.wolflabs.co.uk/IKA_Ultra_Turrax_Tube_Drive.htm).

However I have never personally tried this, nor do I know anyone who has!!

Physcomitrella patens protoplast-PEG transformation 　　　　　　　　　 ver.2.0

Solutions needed for transformation

Driselase/mannitol (for digesting 2platefuls of protonemal tissue)

Dissolve 100mg of Driselase (Sigma D9515) in 10ml of 8% mannitol (not sterile).

* Driselase is a crude enzyme and contains a lot of insoluble materials. Dissolve it in a centrifuge tube and mix the solution occasionally by inverting the tube gently for 15min. Centrifuge the solution at 2500rpm for 5min, then filter-sterilise the supernatant using a 0.22μm filter.

MaMg (for 100ml*)

MgCl₂·6H₂O	3.05g
Mannitol	8g
MES	0.1g

- Dissolve all the ingredients in 90ml dH₂O.

- Adjust the pH to 5.6 with 1M KOH.

- Top up the solution to 100ml.

- Autoclave.

*　 Adjusting pH with a smaller quantity would not be easy. Hence the recipe for 100ml, you will need no more than 10ml per experiment.

- Aliquot the solution in 10ml plastic tubes and store frozen at -20ºC.

PEG-CMS (for 10ml)

Ca(NO₃)₂·4H₂O	0.236g
HEPES	0.0476g
Mannitol	0.728g
PEG	4g

- Dissolve all but PEG in dH₂O in the order shown above. Make sure each chemical is completely dissolved before adding the next one. Bring up the volume to 6ml.

- The pH should be around 7.5 (between pH7 and 8 is acceptable). Use pH paper and 1M KOH (need 30-40 μl) to adjust the pH.

- Add PEG and incubate the solution in the 37°C water bath to dissolve it. Shake it occasionally.

- Top up to 10ml and mix it thoroughly. (The solution is very viscous.). Leave it for several hours at room temperature to stabilise the pH.

- Filter sterilise it and aliquot 1ml each in 1.5ml microtubes. These are stored frozen at －20ºC.

PRM-L (liquid protoplast regeneration medium, for 20ml)

PRM-L = BCD + 5mM NH₄+ 6% (w/v) mannitol + 10mM CaCl₂

Stock B (100´)　　　　　　　　　　　　　　　　　　 200μl

Stock C (100´)　　　　　　　　　　　　　　　　　　 200μl

Stock D (100´)　　　　　　　　　　　　　　　　　　 200μl

0.5M ammonium tartrate (100´)　　　　 200μl (or just add 18.4mg of powder)

TES (1000´)　　　　　　　　　　　　　　　　　　　　 20μl

mannitol　　　　　　　　　　　　　　　　　　　　　　　　 1.2g

1M CaCl₂　　　　　　　　　　　　　　　　　　　　　　 200μl

- Top up to 20ml. Autoclave or filter-sterilise. - If you autoclave, add sterile CaCl₂after autoclaving.

PRM-B (protoplast regeneration medium-bottom layer, for 500ml)

Stock B (100´)　　　　　　　　　　　　　　　　　　　　 5ml

Stock C (100´)　　　　　　　　　　　　　　　　　　　　 5ml

Stock D (100´)　　　　　　　　　　　　　　　　　　　　 5ml

0.5M ammonium tartrate (100x)　　　　　　 5ml (or just add 460mg of powder)

TES (1000´)　　　　　　　　　　　　　　　　　　　　　 0.5ml

mannitol　　　　　　　　　　　　　　　　　　　　　　　 30g

agar (Plant Agar, Duchefa)　　　　　　　　　　 2.75g (0.55%)

- Top up to 495ml and sterilise by autoclaving.

Now add sterile CaCl₂

1M CaCl₂　　　5ml (after autoclaving)

This is used to pour PRM-B plates the day before transformation is carried out. Prepare 3 PRM-B plates per transformation, with roughly 20-25ml PRM-B per plate. These plates are overlain with cellophanes once they are set.

PRM-T (protoplast regeneration medium-top layer, for 100ml)

Stock B (100´)　　　　　　　　　　　　　　　　　　　　 1ml

Stock C (100´)　　　　　　　　　　　　　　　　　　　　 1ml

Stock D (100´)　　　　　　　　　　　　　　　　　　　　 1ml

0.5M ammonium tartrate (100´)　　　　　　 1ml (or just add 92mg of powder)

TES (1000x)　　　　　　　　　　　　　　　　　　　　　 0.1ml

mannitol　　　　　　　　　　　　　　　　　　　　　　　　 6g

agar (Plant Agar, Duchefa)　　　　　　　　　　 0.4g (0.4%)

- Top up to 99ml and sterilise by autoclaving

Now add sterile CaCl₂:

1M CaCl₂1ml (after autoclaving)

- Divide into 10 ml aliquots in sterile Universal tubes.

Note: Protoplast regeneration requires a higher calcium concentration (10mM) than normal growth (1mM).

Transformation protocol

Five days before:

· Prepare fresh homogenate plate subcultures of Physcomitrella on BCD-NH₄ cellophane overlaid plates.

One day before:

· Check the subcultured tissue is not contaminated.

· Pour PRM-B plates and overlay with cellophane.

On the day:

· Defrost PEG-CMS by placing the tubes in warm water. Vortex and pulse-spin to make sure that no precipitate is visible. Make n ´ 300μl aliquots in 10ml centrifuge tubes (where n = No. of transformations). Centrifuge the tubes briefly if the solution is splashed on the side of the tubes.

· Defrost MaMg in warm water. Vortex to make sure that no precipitate is visible. Spin down briefly.

· Aliquot 10-15μg of each DNA in no more than 30μl sterile dH₂O into n ´ 1.5ml microcentrifuge tubes.

· Set a water bath at 45°C.

· Make PRM-L. Add CaCl₂ to 10mM and filter-sterilize.

<Transformation>

1. Scrape 5-day old protonemal tissue from a cellophane-grown culture with a spatula and transfer it into Driselase/mannitol solution. You’ll need 5ml solution per plate of tissue.

NB. 5-6 days old tissue is ideal.

2. Leave it at room temperature (ca 22-25°C) with occasional gentle stirring until filamentous tissue becomes invisible. It takes about 45min – 1hr 30min for well-growing 5 days old tissue.

<<< Every step must be carried out very, very gently from now on! >>>　

3. Carefully transfer the protoplast suspension onto a 100μm filter (nylon or steel mesh) to filter the protoplasts into a sterile conical flask. - Flask, funnel and filter can be set up in advance, wrapped in aluminium foil and autoclaved as a "filter system" The protoplasts will pass through the filter while most of the undigested tissue remains on it.

4. 　Transfer the filtered protoplast suspension to a 10ml centrifuge tube.

5. Spin it down at 800rpm for 4min with the centrifuge brake OFF.

6. Carefully aspirate the supernatant, leaving a very small volume (ca. 0.1-0.2ml) of the supernatant over the (sloppy) pellet. Resuspend the protoplasts initially by gently rocking the tube to loosen the pellet in the residual supernatant, then slowly add 10ml 8% mannitol, mixing this by gently rocking the tube.

7. Spin it down at 800rpm for 4min with the centrifuge brake OFF.

8. Carefully aspirate the supernatant, leaving a very small volume (ca. 0.1-0.2ml) of the supernatant over the (sloppy) pellet. Resuspend the protoplasts initially by gently rocking the tube to loosen the pellet in the residual supernatant, then slowly add 10ml 8% mannitol, mixing this by gently rocking the tube.

9. Set aside a small aliquot (ca. 0.2-0.3ml) of the protoplast suspension to count the protoplast density while recovering the remaining protoplasts by recentrifugation at 800rpm for 4min with the centrifuge brake OFF.

10. Measuring protoplast density: do this during the final centrifugation.

Count 8´1mm² squares (shown in red) and calculate the mean number of protoplasts per 1mm².

Example:　 1mm² square holds 1´10^-4ml (or 0.1μl. The depth of the haemocytometer is 0.1mm). If the mean is 50, the density of the protoplasts in 10ml mannitol is 50´10⁴/ml (5´10⁶ protoplasts in total).

11. Resuspend the protoplasts in the final pellet using MaMg to give an estimated protoplast density of 1.6´10⁶/ml.

Example: If the protoplast count gave a total number of 5´10^6,the protoplasts should be resuspended in (5´10⁶/1.6´10⁶) = 3.1ml MaMg.

12. (optional) Transfer the resuspended protoplasts from the 10ml tube to a wide-topped, shallower container (eg. a Universal bottle or a small Petri dish etc).

NB. This step is to make aliquotting protoplasts easier with a blue tip while reducing the risk of contamination in the next step.

13. Pipette 300μl each of the protoplast suspension with wide-bored blue tip into the 1.5ml microtubes containing 30μl DNA solution.

14. Using a Pasteur pipette, transfer the mixture of protoplasts and DNA to 10ml tube containing 300μl PEG-CMS. Mix gently but thoroughly by stirring the mixture with the tip of the pipette, and by gently sucking up and expelling the whole mixture once.

15. Heat-shock the protoplasts by placing the tubes in the water bath at 45°C for 5 min exactly (NOT LONGER!).

16. Take the tubes out from the water bath and cool them in water at room temperature (22-25°C) for another 10min.

17. Dilute the PEG-protoplast mixture with 8% mannitol over the next 30min-1hr in 6-7 steps. Do not rush this. Wait >3 minutes between each addition to give the protoplasts enough time to recover. The aim is to slowly reduce the concentration of PEG to which the protoplasts are exposed.

Example: Start with 2 successive additions of only 300μl mannitol, followed by two successive additions of 600μl ´ mannitol, followed by two successive additions of 1ml mannitol before finally adjusting the volume to ca. 10ml.

In each dilution step, the protoplasts must be thoroughly, but very gently, mixed with diluents by tilting and/or rolling the tubes (DO NOT SHAKE!).

18. Spin down the protoplasts at 800rpm for 4min with the centrifuge brake OFF. Alternatively, leave tube conatining the diluted protoplasts standing upright on the bench for a few hours. This will allow them to settle slowly.

19. Carefully remove the supernatant - the pellet is very sloppy - and gently add 2ml PRM-L to each tube. Mix gently to resuspend the protoplasts (DO NOT SHAKE!).

20. Place all the tubes in a rack, in a light-tight box and incubate them at 25°C in the dark overnight.

21. Next morning, take the tubes out of the box and incubate under the normal growth-room lights until the PRM-T is ready.

22. Divide the PRM-T in 10 ml aliquots sterile Universal tubes and cool them down by letting them stand in a water bath set at 45ºC.

23. Recover the protoplasts by centriugation at 800rpm for 3 min (Centrifuge brake OFF).

24. Take one of the PRM-T aliquots from the waterbath and transfer it to the sterile flow cabinet. Now uncap the tube.

25. Carefully aspirate the supernatant from the first protoplast tube, quickly add the PRM-T mix, and pour this on 3´ PRM-B plates.

Selection of transformants

After 5 days on PRMB medium the protoplasts can be transferred to selection medium. Simply lift the cellophane overlay from the PRM-B plate and transfer it to a plate containing BCD-NH₄ medium containing the appropriate antibiotic: G418 (30 mg/litre) or hygromycin (20mg/litre).

Resistant colonies do not necessarily represent stably transformed plants! – Physcomitrella can be “unstably” transformed by DNA that is replicated, but not integrated into the genome, so long as selection is maintained. Therefore, it is necessary to go through repeated rounds of selection.

After 2 weeks of initial selection, hand-pick explants of each resistant plant onto BCD-NH₄ medium lacking antibiotic. Allow these to grow for a further 2 weeks.

At the end of this period, re-inoculate a small explant of each candidate plant onto medium once more supplemented with antibiotic. Two weeks of growth on this medium will select those plants that are stably transformed by DNA integrated into the genome.

Integration of DNA is favoured by transformation with DNA as linear fragments. Delivery of circular DNA generates predominantly unstable transformants.

Nucleic acid isolation from moss protonemata

1.　 For all nucleic acid extractions, the following points are important in obtaining good results.

(i) Use protonemal tissue that has been subcultured relatively recently. We routinely isolate nucleic acids from cellophane-grown tissue that is no more than 1-week post-subculture in age. Use of older tissue, or tissue stored at low temperature for long periods results in co-extraction of unidentified crud (probably carbohydrate and phenolics) which is detrimental to subsequent enzyme reactions.

(ii) After harvesting the tissue (typically by scraping off the cellophane) you will have a mass of sloppy wet green material. It is important to remove as much extraneous liquid as possible. We do this by placing the glob of tissue on a sheet of thick filter paper (such as Whatman 3MM chromatography paper), and overlaying with another sheet. Then press down hard, squeezing liquid out of the tissue. Transfer the squeezed tissue to a dry part of the paper, and repeat this twice (i.e. three times in all). A lot of the crud is squeezed out of the tissue in this way.

(iii) Freeze the squeeze-dried mat of tissue in liquid nitrogen. This can then be stored for long periods at -80 C, or extracted immediately.

2. DNA extraction

Commercially available Plant DNA extraction kits can be used to isolate DNA from protonemal tissue with little difficulty　 - we used the “Nucleon Phytopure Plant DNA extraction kit” to isolate genomic DNA for the construction of our Physcomitrella genomic library.

However, we normally use a modified CTAB extraction method for DNA isolation, which is (i) reliable (ii) quick and (iii) cheap.

Extraction buffer:

0.1M Tris-Cl, pH 8.0

1.42M NaCl

2.0% CTAB

20mM Na₂EDTA

2% PVP-40

autoclave and store at room temperature.

Immediately prior to use, add 7ml b-mercaptoethanol, and 10mg ascorbic acid to 10 ml buffer stock. Prewarm at 65 C.

For small-scale (PCR) isolation

1-5mg tissue can be picked from a spot-inoculum, or from a cellophane culture, with a pair of forceps.

Squeeze-dry this and drop it into a 1.5 ml Eppendorf tube.

Samples are ground to a powder while frozen, using a 4mm-diameter glass rod. This is most easily achieved by placing the Eppendorf tube in a plastic “tube raft” (the type used to float tubes in water-baths), with the bottom of the tube immersed in liquid Nitrogen. We use the expanded-polystyrene tube racks that “Falcon” 50ml tubes are packed in as liquid nitrogen baths – A　 well in these racks can be filled to the brim with liquid N₂, and the tube raft holding the Eppendorf tube placed on top, so that the bottom of the tube is bathed in liquid N₂. The glass rods we use are cut to ca. 10cm in length, and the “business end” is roughened with a carborundum stone. The frozen tissue plugs are relatively easily powdered by about 20-30 seconds of grinding.

(Note: Take care not to snap the glass rod, thereby running the risk of impaling your hand with the broken end: we haven’e ever done this, but it is a potential hazard. More frequent is the liklelihood of blistered fingers, if processing a large number of samples. Wearing gloves reduces this!).

This powder can be stored indefinitely at -80, so long as it is not thawed. When processing multiple samples, it is convenient to proceed to this stage for all the samples, before then adding extraction buffer.

To each 1ml of extraction buffer add 10 ml 10mg/ml Rnase A (pre-boiled for 10 minutes to denature any contaminating DNase). Add 100 ml to the powdered tissue, mix and incubate at 65 C for 5 min.

Add 100 ml chloroform-iso-amyl alcohol (24:1) and mix briefly but thoroughly: the aim should be to avoid mixing so violent as to shear high-molecular-weight DNA.

Separate the phases in a microcentrifuge for 10 min.

Transfer the upper phase to a fresh tube and add 70 ml iso-propanol

Mix well and centrifuge immediately for 5 minutes.

Decant the supernatant and wash the pellet with 70% ethanol. Drain and air-dry the pellet. Dissolve in 15 ml TE buffer (10mMTris-Cl, pH8 - 1mM Na₂EDTA).

This provides good quality DNA readily amplifiable by PCR.

For large-scale (Southern Blot) isolation

The same basic-procedure is used, but scaled-up.

Harvest one plate of tissue (by scraping protonemata from the cellophane) and squeeze-dry.

Freeze the tissue in liquid N₂ and grind with a small mortar and pestle. (The liquid N₂ treatment facilitates effective rupture of the cells. It is not necessary to use a pre-chilled mortar). Add 1ml extraction buffer and continue gentle mixing with the pestle to obtain a smooth paste.

Add a further 1ml extraction buffer, stirring to obtain a uniform homogenate and transfer this to a suitable high-speed centrifuge tube (e.g. a 15ml Corex tube). Wash out residual homogenate into the tube with another 1ml aliquot of buffer.

Add 30 ml 10mg/ml Rnase A and incubate at 65 C for 5 min.

Add 3 ml chloroform/iso-amyl alcohol and vortex to emulsify.

Separate the phases by centrifuging at 10,000rpm for 10 min. in a swingout rotor (e.g.Sorvall HB-4)

Transfer the upper (aqueous) phase to a fresh tube and precipitate the DNA by adding 2.1 ml iso-propanol, mixing well and immediately centrifuging at 10,000 rpm for 5 min in a swingout rotor.

Wash the pellet with 70% ethanol and air-dry. Dissolve the pellet in 2 x 100 ml TE, transferring it to a 1.5ml Eppendorf tube.

Centrifuge this for 2 min at 12,000 x g in a microcentrifuge. You may observe a translucent pellet (carbohydrate). Recover the supernatant carefully and transfer it to a clean tube for storage at -20 C.

Typically, this provides sufficient DNA for at least 5 Southern blots.

Digestion of DNA

Although the design of any particular experiment may require the use of a specific restriction enzyme for Southern blot analysis of Physcomitrella DNA, it should be noted that some enzymes cleave Physcomitrella DNA to a greater extent than others. This probably relates to the extent and distribution of methylation of the moss genome.　 - This has been discussed by Krogan & Ashton (1999) who demonstrated that enzymes with recognition sites subject to C-methylation at CG and CNG sequences were less effective in digesting Physcomitrella DNA.　 In our hands, HindIII routinely yields the most effective digests.

The most heavily methylated sequences in Physcomitrella may lie outside the coding sequences.　 We have noted that in screening a lambda genomic library for numerous Physcomitrella genes, that in every case the cloned sequence was located at the end of the inserted genomic fragment. This implies that the sites most accessible to the restriction enzyme used to generate the cloned fragments (a partial digest using Sau3A) were undermethylated sites within transcribed regions of the genome.

3. RNA extraction

The isolation of RNA from most biological materials is not difficult. Nevertheless, a body of myth has built up that leads some workers to take precautions that lie on the borders between paranoia and superstition. This includes the segregation of laboratory supplies “for RNA extraction only”, the treatment of solutions with diethyl pyrocarbonate and the baking of glassware at temperatures rarely seen outside a thermonuclear explosion. These precautions are largely unnecessary, so long as simple common-sense informs your procedures.

1. There is always more Ribonuclease in the tissue you are extracting than there is ever likely to be in the solutions/chemicals/glassware you use.

2. SDS is very effective at eliminating any trace contamination by ribonuclease than may occur. – For example, I routinely use the same centrifuge tubes used for DNA isolation (during which procedure they contain a suspension supplemented with RNase A at a final concentration of 100μg/ml) as I do for RNA extraction. – Thorough washing of the centrifuge tubes by soaking overnight in 0.1%SDS ensures that I have never suffered from RNase degradation of my samples.

Physcomitrella protonemal tissue presents no great difficulties for the extraction of RNA, so long as the general preliminary procedures used in DNA isolation are followed: namely the “squeeze-drying” of the tissue to remove residual liquid, prior to extraction.

There is little endogenous ribonuclease activity, and the tissue is readily amenable to RNA isolation using most commercially available kits. However, I typically use an aqueous-SDS/phenol extraction procedure that has served me well for the past 30 years.

Extraction buffer:

0.1M Tris-HCl, pH9

0.5% SDS

2% PVP-40

5mM Na₂EDTA

Autoclave and store at room temperature.

Immediately prior to use, add 7ml b-mercaptoethanol to 10ml extraction buffer.

For small scale extractions:

Harvest the tissue by scraping from the cellophane, and squeeze-dry as described for DNA extraction. Tissue equivalent to up to one 7-day old 9 cmhomogenate plate can be processed entirely in 1.5 ml Eppendorf tubes: Here is a relatively high-throughput extraction protocol.

Harvest tissue from cellophane and squeeze-dry to produce a plug of tissue. Put this in a 1.5ml Eppendorf tube, and freeze the sample by dropping it in liquid N₂. At this stage, the tissue can be powdered as described for DNA small-scale preparation (above) and stored at -80 C indefinitely.

Extraction:

To each tube containing powdered tissue, add 500 ml extraction buffer, and 500 ml phenol-chloroform-iso-amyl alcohol (25:24:1), cap the tube and vortex thoroughly to produce an emulsion.

Centrifuge at 12,000 x g for 2 minutes (microcentrifuge), and recover the upper (aqueous) phase, transferring it to a fresh tube containing 50 ml 3M Na-acetate (pH 5.2).

Mix well and place the tubes on ice for 2-5 minutes, then centrifuge at 12,000 X g for 2 minutes.

This is necessary to remove the very large amount of carbohydrate (principally pectin) that co-extracts with the nucleic acids.

A substantial gelatinous precipitate forms. Transfer the supernatants to fresh 1.5ml Eppendorf tubes, add 1ml ethanol, mix and incubate at -20 C for 20 minutes to precipitated RNA and DNA. Centrifuge at 12,000 X g for 5 minutes.

Discard the supernatant, drain the pellet and allow it to air-dry briefly. The pellet contains DNA and RNA.

Dissolve the pellet in 200 ml sterile water, and add 30 mg solid NaCl. Mix well to dissolve the salt completely and incubate at 4 C for at least 4 hr (Overnight incubation is often more convenient)

High-molecular weight RNA (including mRNA) is recovered by selective precipitation with NaCl. At 2.5M NaCl, DNA and low molecular weight RNA remain in solution, high molecular weight RNA species precipitate.

To recover the RNA, centrifuge at 12,000 x g for 10 minutes at 4 C. Carefully aspirate the supernatant using a micropipette. Care is required, since the (RNA) pellet may be relatively sloppy, having been centrifuged through a relatively viscous DNA-rich supernatant.

Resuspend the pellet by vortexing with 200 ml 2.5M NaCl, and recentrifuge for 5 minutes at 12,000 x g. This time, the pellet should pack tightly as residual DNA is washed out.

Discard the supernatant, and wash the pellet with 70% ethanol by resuspension and repelleting at 12,000 x g for 5 minutes. Drain the pellet and allow it to air-dry thoroughly.　 Finally, the pellet can be dissolved in 30 ml sterile water and stored at -20 C.

Typically, approximately 30-50 mg RNA is obtained by this method. Agarose gel electrophoresis reveals it to be substantially composed of cytoplasmic and chloroplast rRNA species (the latter appearing as a ladder of fragments resulting from the “hidden breaks” within the molecules when a denaturing gel is used). Occasionally a trace of residual DNA is apparent in such preparations. If this is a problem, this can be removed by repeating the 2.5M NaCl wash step in the protocol.

Large-scale isolation.

The same basic protocol is followed, but scaled up appropriately.

For 1 or 2 plates of protonemal tissue (7 days post subculture), harvest the tissue and squeeze-dry thoroughly.　 At this stage, tissue can be placed in “envelopes” made by folding aluminium foil, and frozen in liquid N₂, for storage at -80 C indefinitely.

Use a small mortar and pestle. To aid in homogenisation, a small quantity (ca. 100mg) of glass beads (80-100mesh) or sand can be added to the mortar. – It is recommended to (i) acid-wash and (ii) autoclave this material in advance.

Pre-chill the mortar and pestle by pouring a little liquid N₂ into it, then add the lump of squeeze-dried, frozen tissue and grind to a powder.

As the powder thaws, add 1ml extraction buffer and homogenise to a smooth slurry. Transfer this to a centrifuge tube (e.g. 15 ml Corex tube). Wash residual material from the mortar and pestle with a further 1ml extraction buffer, and bulk this with the homogenate.

Add 2 ml phenol-chloroform-iso-amyl alcohol (25:24:1) and vortex thoroughly, to produce an emulsion.

- Speed is the principal criterion to observe: each sample should take no longer than 2 minutes to process to the emulsion stage.

If multiple samples are being processed, this emulsion can be stored on ice prior to preceeding to the subsequent centrifugation step: I generally do not accumulate more than 4 such emulsions at one time.

Clarify the emulsion by centrifuging in a swingout rotor (e.g. Sorvall HB-4 rotor) at 10,000 rpm x 5 min.

Transfer the upper (aqueous) phase to a clean centrifuge tube, and add 0.2 ml 3M Na-acetate, pH 5.2 and 4.5ml ethanol. Mix well and place at -20 C for 30 minutes.

Recover the ethanol-precipitate by centrifugation in the swingout rotor at 10,000 rpm x 5 min. Discard the supernatant. Typically, you will observe a substantial translucent pellet in the bottom of the centrifuge tube. – This contains a substantial quantity of carbohydrate which must be removed. (Note that unlike the miniprep method, it is not so easy to separate the carbohydrate from the supernatant in a larger tube, prior to the ethanol-precipitation step – but you can try it if you like).

Drain the pellets well, and resuspend in 200μl sterile water by vigorous vortexing. Keep the tube on ice at this stage. This produces a very viscous solution which may still contain lumps of pellet. Using an automatic pipette, remove the liquid and transfer it into an Eppendorf tube, held on ice.

Add a further 200μl sterile water to the centrifuge tube, to recover the remaining pelleted material. After vigorous vortexing, the residual material should completely dissolve. Recover this and add it to the first aliquot in the Eppendorf tube. Vortex this vigorously to ensure the pellet has completely dissolved. At this stage, you should have a viscous solution with a volume of approximately 500 μl.

To this, add 25μl 5M NaCl, mix well and incubate on ice for 20 minutes. Centrifuge in a microcentrifuge (12,000 x g , 5 min) to pellet the carbohydrate, which forms a substantial, compact gelatinous pellet.

Transfer the supernatant to a clean Eppendorf tube, and add 75 mg NaCl. Vortex thoroughly to dissolve the NaCl, and incubate overnight at 4 C. – It is often convenient to prepare a series of tubes containing 75mg aliquots of solid NaCl to which the supernatants can be added.

Pellet RNA by centrifugation in a microcentrifuge at 12,000 x g, 10 min, 4 C. Remove the supernatant and wash the pellet by vigorous resuspension in 300μl 2.5M NaCl. Immediately centrifuge at 12,000 x g, 10 min, 4 C.

Finally, wash the pellet by vigorous resuspension in 200μl 70% ethanol, and centrifugation in the microcentrifuge. Drain the pellet (carefully – it may not adhere firmly to the wall of the centrifuge tube!!!) and air-dry until all traces of ethanol have evaporated. The pellet can be dissolved in an appropriate volume (ca. 50μl) of sterile water for storage at -20 C.

RNA obtained by this procedure can be used directly for Northern blot analysis, or for the subsequent enrichment of poly(A)-containing mRNA, by oligo-dT affinity chromatography.

We routinely use RNA prepared in this way for translation, in vitro, for the synthesis of cDNA in the construction of cDNA libraries and the labelling of probes for hybridisation with microarray chips.

Quality control

The yield and integrity of the RNA isolated can be determined by measuring tha absorbance at 260nm, and by agarose gel electrophoresis. RNA can be electrophoretically analysed in the same way as DNA, in a 1.4% agarose minigel buffered with Tris-Borate-EDTA, followed by staining with ethidium bromide.

Typically, you should observe a number of discrete bands that correspond to the principal cytosolic rRNA species and the 16S chloroplast rRNA. You may also observe denaturation products of 23S chloroplast rRNA (Note: this chloroplast rRNA naturally contains “hidden breaks” – specific sites in the chain are cleaved, in vivo. The secondary structure of rRNA maintains the integrity of the molecule in the ribosome, but these breaks become apparent upon extraction of the RNA. These smaller bands are therefore NOT indicative of degradation during the extraction process). Degraded RNA typically appears as a background “smear” along the length of the gel. If you observe such a smear, you should determine whether it results from degradation during extraction, or degradation during electrophoresis: the commonest use of agarose minigel tanks is to analyse plasmid DNA isolated from bacterial cultures. Most plasmid DNA isolation procedures utilise a ribonuclease digestion stage, and it is not uncommon for RNase to contaminate gel tanks and trays. This is one instance where it is important to ensure that your equipment is RNase-free, since purified RNA is very susceptible to degradation. Simple washing of the gel tank in 0.1% SDS is sufficient to ensure that RNase degradation during electrophoresis does not occur.

Example of RNA electropherogram:

Each well contains 1/15 of the RNA isolated from a half-plate of protonemal tissue. Note the 5^th lane in the top tier of samples. This RNA sample appears degraded relative to the others, with weaker staining of the principal bands, and a diffuse, but obvious, lower molecular weight smear.

Southern blot hybridisation of genomic DNA

Digest genomic DNA (2.5 - 5μg for Physcomitrella) with the appropriate restriction enzymes and check a small aliquot on a TBE minigel to ensure complete digestion.

Gel electrophoresis

10x Tris-borate-EDTA ("TBE") Buffer:

Tris-base　　　　　　　　　　　　　　　　 108g

Boric Acid　　　　　　　　　　　　　　　 55g

Na₂EDTA　　　　　　　　　　　　　　　　 9.3g

H₂O　　　　　　　　　　　　　　　　　　　 to 1 litre

pH should be about 8.3. Store in 500ml aliquots in 500ml bottles

10x TBE buffer will precipitate out over time, once a bottle is opened and the solution exposed to air. Generally, we prepare a litre of stock, and immediately use 500ml to make 5 litres of 1x "working-strength" TBE buffer by diluting it with double-distilled water. The remaining 500ml is kept unopened until it is time to prepare another 5 litres of 1x TBE.

To prepare a gel, weigh out the appropriate mass of agarose to be dissolved in the desired volume of 1x TBE.

Minigels are typically 50ml in volume. The BioRad midi-gel uses 100 to 150ml (as required: a deeper gel allows a larger volume of sample to be loaded into the wells)

Gel concentration is important: the concentration should be chosen to optimise resolution of the desired DNA fragments.

Recommendations:

Fragment size　　　　　　　　　　　　 Gel concentration

200bp - 2kb　　　　　　　　　　　　　　　　　　　　 1.5%

400bp - 3kb　　　　　　　　　　　　　　　　　　　　 1.2%

500bp - 5kb　　　　　　　　　　　　　　　　　　　　 1%

800bp - 8kb　　　　　　　　　　　　　　　　　　　　 0.8%

1kb - 10kb　　　　　　　　　　　　　　　　　　　　　 0.7%

For genomic digests used for Southern blotting, I usually use 0.6 to 0.7% agarose gels.

TBE minigels can also be used to assess the recovery and quality of RNA. In this case, a 1.2-1.5% gel is recommended. Additionally, the gel tank should be RNase-free. (See "RNA extraction") for an example.

For a 50 ml gel, put the agarose in a 250ml conical flask and add the 1x TBE. Do not swirl this suspension!! - this will result in lumps of unmelted agarose being deposited up the walls of the flask.

Melt the agarose in the microwave oven.

Guideline: 50 ml of a 1% gel requires 2 minutes at power level 5

Check that the agarose has completely dissolved by gently swirling the conical flask

(Danger: this will be VERY HOT!!!!)

Now allow the agarose solution to cool until the flask is hand-hot. At this point, Ethidium bromide (EtBr) can be added to the gel. This is routine for mini-gels, but not recommended for gels where Genomic DNA is being used for Southern Blot analysis: ethidium bromide binds DNA by intrcalating between the base-pairs, and it affects the mobility of the DNA. If accurate size-determination is necessary, run the gel without EtBr, and stain the gel after running. - This will take longer, but is more accurate. However, for many purposes (e.g. checking plasmids, checking digests, etc. the EtBr can be added to the gel before pouring: add 1μl 10mg/ml EtBr per 10 ml gel solution.

Safety note: Ethidium bromide is a dangerous mutagen: wear gloves and handle with care.

When the gel has set, remove the comb and tape (BioRad) or end-plates (FlowGen), and submerge in 1x TBE electrophoresis buffer. The BioRad mini-gel tanks take ca. 250 ml buffer. The FlowGen flat-bed tank takes 50 ml.

To each DNA sample (e.g. the the RE digestion mixture) add a ¹/₁₀ volume of 10x gel loading buffer:

10x DNA gel loading buffer:

125mM EDTA - 50% glycerol-Bromophenol Blue to colour

Stored in aliquots at room temperature.

(note: Xylene cyanol can also be included as a second tracking dye)

Run the gel until the BPB dye reaches the end of the gel. The most accurate size determination requires slow running (ca. 5V/cm voltage gradient). In fact, for most purposes the gel can be run faster than this. For the BioRad mini and midi gel tanks (the same length), run at 100V constant voltage; for the FlowGen tank, 60V can be used. However, while this is adequate for most "quick & dirty" applications (checking digests etc., I do not recommend it for genomic Southern blots where undistorted banding patterns are desired.

For Genomic Southern blots, I strongly recommend using the BioRad tanks. The greater volume of electrophoresis buffer means that the gel does not become too hot, which can distort the bands. Do not run the gel at a voltage greater than 50V (21mA), which results in migration to the end of the gel in about 4 hours. If you have time, run the gel slower!!

After running

Stain the gel in 1μg/ml EtBr solution for 15 minutes (if it isn't already in the gel), then destain in ddH₂O for 10 minutes. Photograph the gel on the UV-light box using a fluorescent ruler alongside the Mr marker track.

Mr Markers

There are numerous proprietary Mr marker sets available. It is, however usually considerably more economical to prepare your own. We use bacteriophage lambda DNA digested with either HindIII or PstI

Blotting

We use semidry alkaline transfer:

1. Transfer the gel, upside-down, to a suitable container (pyrex baking dish or pippette tip box lid, depending on size) and wash with 0.25M HCl for 15 minutes only, using the shaking platform at low sppeed to ensure the gel is washed thoroughly.

- inverting the gel ensures that the filter will be placed on the flat side of the gel, rather than the concave "meniscus-side". The HCl partially depurinates the DNA, fragmenting the larger fragments, in situ and ensuring that the HMW fragments are transfered out of the gel more efficiently during blotting. It is important not to exceed the 15 minutes, or the DNA will become too degraded.

2. Rinse the gel twice with dd H₂O

3. Wash the gel in 0.4M NaOH, twice, for 15 minutes each time.

4. Remove all the liquid. Carefully overlay the gel with a sheet of Biodyne B membrane, cut to the size of the area containing the DNA. Ensure no air-bubbles are trapped between filter and gel.

5. Now overlay the filter with a few sheets of 3MM paper cut to the same size as the filter. Trim away any excess gel and discard it. Pile up a stack of paper towels on the 3MM paper and place a suitable weight on the top to compress the stack.

Blotting can proceed overnight, if desired, but will be complete sooner. A few hours is enough to complete transfer of the DNA. This method uses the liquid in the gel to transfer the DNA onto the filter, and is much easier than messing about with wicks and reservoirs.

Biodyne B is a positively-charged nylon membrane. Under alkaline conditions the DNA is simultaneously denatured and covalently bound to the membrane. Do not used Biodyne A, which is uncharged. We also UV-crosslink the DNA to the membrane in a "belt & braces" approach.

After blotting.

Dismantle the blot and rinse the filter twice with 5x SSC (about 3 minutes per wash).

Air-dry the filter and bind the DNA to it by using the Stratalinker set on "autocrosslink". - Place the filter, DNA side up, on a piece of 3MM paper. Put this into the Stratalinker and press "Autocrosslink". The LED display should read "1200".

Press "start". The machine will bleep when it is finished (about 30 sec).

The blot can now be stored, wrapped in Saran Wrap, or hybridised immediately.

Hybridisation:

2x Hybrid buffer stock:

1M NaPO₄ pH 6.4　　　　　　　　　　　　　　　　　　　　　 20ml

20x SSC　　　　　　　　　　　　　　　　　　　　　　　　　　　　 100ml

200x Denhardts (no BSA)　　　　　　　　　　　　　　　 10ml

Dextran sulphate　　　　　　　　　　　　　　　　　　　　　 20g

H₂O　　　　　　　　　　　　　　　　　　　　　　　　　　　　　　　 to 170ml

stir until dissolved on stirring hot-plate, then add

10% SDS　　　　　　　　　　　　　　　　　　　　　　　　　　　　　　　　　 20ml

10mg/ml sonicated, denatured calf-thymus DNA　　　 10ml

Store at -20 C

"SSC" is "standard saline citrate: 20x SSC = 3M NaCl-0.3M Na citrate

NaPO₄ pH 6.4 is prepared by mixing 73.5ml 1M NaH₂PO₄and 26.5ml 1M Na₂HPO₄

200x Denhardts contains 4% (w/v) Ficoll-400 and 4% (w/v) PVP-40: store aliquots at -20 C

ctDNA is sonicated until sheared to less than 1kb average length, then boiled, snap-cooled and stored at -20 C

For Southern blot hybridisation/prehybridisation, mix equal volumes of 2x hybridisation buffer and ddH₂O (for a 9cm x 6.5cm filter, a total volume of 5ml is sufficient).

Place the filter in a Hybaid rotisserie bottle with hybridisation fluid and prehybridise at 65 C for at least 2 hours.

Hybridise by adding denatured, labelled probe DNA to the bottle and continuing the incubation in the Hybaid oven overnight.

Washing

Wash the filter to remove unhybridised probe. Use different salt concentrations and temperatures to detect identical and non-identical but similar (paralogous) genes. High stringency washing for perfectly matched hybrids is with 2x SSC-0.1% SDS twice, followed by 0.1x SSC-0.1% SDS twice, at 65 C for at least 15 minutes per wash. However, lower stringencies might use only 2x SSC, or 5x SSC, and lower wash temperatures.

Remove the filter and wrap it in Saran Wrap (don't let it dry!!). Stick this to a piece of old X-ray film with a radioactive ink marker label, and expose to X-ray film at -70 C, in a cassette containing an intensifying screen for as long as necessary. - Monitor the filter with the beta-monitor to gauge how strong the radioactive signal is.

MOPS-Formaldehyde denaturing gel for RNA blots

Use a gel tank and comb pre-washed with 0.1% SDS and thoroughly rinsed with dH₂O, to ensure an RNase-free environment. Ideally, use a designated "RNA-only" gel tank.

10x MOPS buffer:

MOPS　　　　　　　　　　　　 0.2M　　　　　　　　　　　　　 20.94g

Na Acetate　　　　　　　　 50mM　　　　　　　　　　　　 8.4 ml 3M NaAc pH7.0

EDTA　　　　　　　　　　　　　 10mM 　　　　　　　　　　　 10ml 0.5M Na₂EDTA pH 8.0

pH　　　　　　　　　　　　　　　 7.0　　　　　　　　　　　　　　　 with NaOH

　　　　　　　　　　　　　　　　　　　　　　　　　　　　　 H₂O to 500ml

NB: Prepare the NaOAc and EDTA solutions and autoclave them. Add them to solid MOPS and dilute with dd water. Adjust to pH 7.0 with 5M NaOH and make up to the final volume. This solution goes off if stored at room temperature, so store in 250ml aliquots in polypropylene bottles at -20 C.

Prepare gel:

For 　　　　　　　　　　　　　　 50ml　　　　　　　 100ml　　　　　　 150ml

Agarose　　　　　　　　　　　 0.7g　　　　　　　　 1.4g　　　　　　　　 2.1g

ddH₂O　　　　　　　　　　　　 42.3ml　　　　　　 84.7ml　　　　　　 126.9ml

Dissolve agarose and cool, then add

10x MOPS buffer　　　　 5ml　　　　　　　　 10ml　　　　　　　 15ml

40% Formaldehyde　　 2.7ml　　　　　　　 5.3ml　　　　　　　 8.1ml

Note: Formaldehyde is noxious: wear gloves and handle only in fume hood!!

This protocol is modified from Davis (1986), but uses lower concentrations of formaldehyde. It remains very effective.

Pour gel into tray and allow to set. When set, remove comb and place gel in tank with 1x MOPS buffer as running buffer (using ddH₂O to dilute). Add the running buffer so that the gel is not quite submerged: the buffer level should be just below the level of the top of the gel.

RNA loading buffer:

Deionised formamide　　　　　　 7.2ml

10x MOPS　　　　　　　　　　　　　　　　　　　　　 1.6ml

37% Formaldehyde　　　　　　　　　　　　　　 2.6ml

Glycerol　　　　　　　　　　　　　　　　　　　　　　 1ml

H₂O　　　　　　　　　　　　　　　　　　　　　　　　　　 1.8ml

Bromophenol Blue　　　　　　　　　　　　　　 0.8ml saturated solution

Store at -20 C in 1ml aliquots.

NB: Deionised formamide is prepared as follows: Pour 100ml formamide into a glass beaker and add a spoonula-full of ion-exchange resin "AG501-X" (BioRad), Amberlite MB-1" or "Amberlite IRN-150L" (BDH). Stir on a magnetic stirref for about 30 minutes and filter through Whanman No. 1 paper. This should be used within 1 week if stored at room temperature, but can be regenerated by repeating this procedure.

Calculate the volume to be loaded in each track (5-15μg total RNA:　 less if poly(A)⁺RNA), and ensure equal loading in each track. This can be based on (i) UV absorbtion measurements (A₂₆₀) combined with (ii) estimates from the staining intensity of samples on TBE minigels. Mix the RNA with an equal volume of loading buffer (mixing with 2 volumes is better, but not always practical).

Heat at 80 C for 2 minutes and snap-cool, on iced water. Dry-load the slots of the gel and apply current.

For the Bio-Rad mini and midi gel tanks, run at 50V (approx 9mA) for 5-10 minutes to run the sample into the gel - the BPB should migrate about 5mm. Top up the tank with buffer to submerge the gel, and continue running until the BPB reaches the end of the gel (approx. 4 to 5 hr)

Northern blotting

Prepare a stack of 3MM paper rectangles the same size as the area of the gel to be blotted.

Soak each in 20x SSC, building up a stack in a plastic box containing the 20x SSC reservoir. (Pippette tip box lids are a convenient size.)

Place the gel, upside-down, onto the stack of papers, and carefully overlay with a sheet of Biodyne B membrane cut to the size of the area to be blotted. - Mark the positions of the tracks with a pencil (NOT ink!!), beforehand. Overlay this with several sheets of 3MM paper of the same size, and trim away any overhanging bits of gel.

Place a stack of paper towels cut to the correct size on top of this and place a weight on the stack (a small retort-stand base, an aluminium tube block or similar).

Blot overnight at rt.

After blotting:

1. Wash the filter briefly in 6x SSC

2. Air-dry the filter then fix the RNA by baking at 80 C for 2 hours.

3. Wash the filter in 5% acetic acid for 5-10 minutes

4. Stain the filter by immersing it in 0.5M Na Acetate, pH5.2 - 0.04% Methylene Blue for 30-60 seconds (ca. 10 ml, shake to ensure even coverage)

5. Wash the filter with several washes of dd water, until the RNA bands are clear against a white background.

7. Photograph the filter next to a ruler, then wash out the stain by gentle shaking in 5% SDS

8. Rinse briefly with dd water, then the filter is ready for prehybridisation.

Hybridisation:

For Northern blot hybridisation/prehybridisation, mix equal volumes of 2x hybridisation buffer and deionised formamide (for a 9cm x 6.5cm filter, a total volume of 5ml is sufficient).

Place the filter in a Hybaid rotisserie bottle with hybridisation fluid and prehybridise at 42 C for at least 2 hours.

Hybridise by adding denatured, labelled probe DNA to the bottle and continuing the incubation in the Hybaid oven overnight.

Washing

Wash the filter with 2x SSC-0.1% SDS twice, followed by 0.1x SSC-0.1% SDS twice, at 50 C for at least 15 minutes per wash.

Plasmid DNA miniprep: I. Mini-boiling method

From Del Sal et al (1998): Nucleic Acids Res. 16: 9788. Adapted from Holmes & Quigley (1981) Anal. Biochem. 114: 193-197

This method produces high-quality plasmid DNA suitable for all subsequent procedures. However, it is not suitable for all E. coli strains. Some contain DNase that is not removed by this method (HB101 is the principal offender). Nevertheless, this method is good for XL-1 Blue, DH5α, DH10b and most other strains that are commonly used for cloning. Strains that are recalcitrant can be recognised when you try to digest the plasmid with restriction enzymes: instead of getting clean bands, you get a fuzzy smear. If　 your strain is not suitable for this method, use the alkaline lysis method, instead.

Solutions:

STET buffer:

8% (w/v) sucrose

50mM Tris-Cl pH 8

50mM Na₂EDTA pH 8

0.1%(v/v) Triton X-100

Autoclave and store at r.t.

5% (w/v) CTAB

(= cetyltrimethylammonium bromide = hexadexcyltrimethylammonium bromide)

Store at r.t. - on cool days, CTAB can precipitate out. If this happens, it is easily redissolved by placing the bottle in a hot water bath for a few minutes.

1.2M NaCl

(Prepare by mixing 12 ml 2M NaCl and 8ml sddH₂O: store at r.t.)

3M Na Acetate pH 5.5

Procedure

Inoculate 4ml cultures of bacteria in Sterilin Bijou bottles (LB + antibiotic) and shake overnight at 37 C.

Next morning, freshly prepare STET plus lysozyme at 2mg/ml. (= STETL). Lysozyme powder is stored at -20 C, so allow the bottle to equilibrate to room temperature before opening. This prevents condensation forming on the powdered enzyme. Prepare only enough for use.

Prepare a 1.5ml Eppendorf tube for each culture. Harvest 3ml from each culture (fill the tube, spin down the bacteria for 1 min in the microfuge at full speed and decant the supernatant. Top up the tube with more culture and repeat).

Drain the pellet well. Resuspend each pellet in 200μl STETL by vortexing or tube strumming. Ensure that the pellet is completely resuspended. Incubate at room temperature for 5-10 minutes.

Transfer the tubes to a vigorously boiling water bath. It is very important that the tubes are in direct contact with the water (i.e. NOT in a heating block). Incubate in the boiling water for EXACTLY 45 seconds (not more!!!). The suspension should turn white and opaque.

Spin the tubes in the microfuge at full speed for 10 minutes. This produces a loose, slimy white pellet comprising denatured chromosomal DNA and protein. Fish this out with a toothpic and discard. Retain the supernatant.

To the supernatant, ass 8μl CTAB and mix. You should immediately see a precipitate forming. Now, immediately spin the tubes in the microfuge for 5 minutes.

After spinning, you should see a fibrous white pellet in each tube. Remove the supernatant completely (I use a drawn-out pasteur pipette attached to a vacuum line).

Dissolve the pellet in 300μl 1.2M NaCl The pellet is very difficult to dissolve. Vortexing is not usually sufficient. After initial vortexing, vigorous strumming of the tube on a wire-rack should completely dissolve the pellet. - Because the tube contains residual CTAB (a detergent) this also causes extensive foaming.

When the pellet has dissolved, precipitate the DNA by adding 750μl EtOH: incubate at -20 C for 20-30 minutes and recover the DNA by spinning at full speed in a microfuge for 5 min.

Drain the pellet well.

Note: Del Sal et al claim that at this point, the DNA can be washed and dissolved in TE and is pure enough for sequence analysis. However, the pellet contains lots of RNA, and I prefer to clean up the DNA further.

Prepare a solution containing 10mMTris-Cl - 1mM Na₂EDTA - 20μg/ml RNase A. ("TERNase"): This is most easily done by mixing T₁₀E₁ with 1 μl 10mg/ml RNaseA per ml of TE. Commercial preparations of RNase usually contain some contaminating DNase, so it is important to pretreat RNase stock solutions by boiling them for 10 minutes (!!). I usually prepare a 1ml stock solution of 100mg/ml RNaseA in 50% glycerol, in a screw-cap Eppendorf tube, which is then placed in a boiling water bath for 10 minutes. This can be used to prepare a diluted 10mg.ml stock solution, also in 50% glycerol. these are stored at -20 C.

BE CAREFUL WHEN PREPARING RNase SOLUTIONS: WE CARRY OUT A LOT OF RNA EXTRACTIONS, SO TAKE CARE TO AVOID SPILLS. (The reason I stipulate screw-cap tubes for RNase stocks is to avoid aerosol sprays on opening flip-top tubes)

Dissolve the DNA pellet in 200μl TERNase and incubate at 37 C for 15-20 minutes.

Phenol-chloroform extract with 200μl phenol-chloro. (Vortex, spin 2 min in microfuge at full speed). Transfer the (upper) aqueous phase to a fresh tube taking care not to transfer the organic/interphase material.

Add 20μl 3M NaAc and 450μl EtOH, mix and precipitate the DNA at -20 C for 20-30 min.

Spin down (5 min, full speed in microfuge) and discard the ethanol. Wash the pellet by vortexing with 200μl 80% EtOH and re-pellet (5min at full speed in microfuge).

Drain the pellets well and dry the DNA completely by standing inverted on tissue paper.

When the DNA is completely dry, dissolve the pellet in 30μl T₁₀E₁ and store at -20 C.

Plasmid produced in this way (expect 100-200 ng/μl) is very clean and amenable to all subsequent procedures (digestion, ligation, sequencing) without further treatment.

Plasmid DNA miniprep: II. Alkaline lysis method

From Birnboim & Doly (1979) Nucleic Acids Res. 7: 1513-1523. This method is the basis of most commercial plasmid miniprep kits (e.g. Quiagen).

Solutions:

GTE:

50mM Glucose

10mM Na₂EDTA pH8

25mM Tris-Cl pH8

Autoclave and store at room temperature.

Alkaline SDS

0.2M NaOH

1%(w/v) SDS

Prepare freshly from concentrated stocks of NaOH and 10% SDS each time.

3M Na Acetate pH 4.8

Filter-sterilise and store at room temperature.

Procedure:

Grow 4ml miniprep cultures in Sterilin Bijou bottles containing LB + antibiotic, and harvest 3ml aliquots in 1.5ml Eppendorf tubes as described for the mini-boiling method.

Dissolve lysozyme to 2mg/ml in GTE (prepare freshly just before use) and resuspend the bacterial pellets in 100μl of this solution.

Incubate on ice for 30 minutes.

Add 200μl alkaline SDS and gently mix by inverting the tube. The suspension should necome clear an viscous. Incubate on ice for 5 minutes.

Add 150μl 3M NaAc pH 4.8 and mix well, but gently, by inverting the tube. Incubate on ice for 60 min.

Spin at full speed in a microfuge for 5 minutes. A substantial white precipitate (chromosomal DNA, SDS-protein complex) will pellet along the rear wall of the tube.

Carefully remove 400μl supernatant with a blue Gilson tip and transfer this to a fresh tube. Add 1ml EtOH and mix to precipitate the plasmid DNA (20-30 min at -20 C).

Recover the DNA by spinning at full speed in the microfuge for 5 min. Drain the pellet well.

Dissolve the pellet in 200μl TERNase and incubate at 37 C for 15 min.

Extract with 200μl phenol-chloroform, and recover the aqueous phase.

Ethanol-precipitate the DNA by addition of 20μl 3M NaAc and 450μl EtOH (mix well and incubate 20-30 min, -20 C).

Spin down (5min, full speed in microfuge) and discard the supernatant. Wash the pellet by vortexing in 200μl 80% EtOH and spin for 5 min at full speed in the microfuge.

Drain the pellet well and dry completely. Dissolve in 30μl T₁₀E₁ and store at -20 C.

Radiolabelling of DNA with α-³²P-dNTPs

We use the "oligolabelling" method of Feinberg & Vogelstein (1983): Analytical Biochemistry 132: 6-13

Stock solutions:

1M HEPES-KOH pH6.0

"TM": 0.25M Tris-Cl, pH 8

　　　　　 25mM MgCl₂

　　　　　 50Mm β-mercaptoethanol

dNTP stocks:　 20mM each dNTP

"OL":　 90 A₂₆₀/ml random hexanucleotides (Pharmacia pd(N)₆ dissolved in 　　　　　　　　　　　 1mM Tris-Cl pH 7.5 - 1mM Na₂EDTA)

***************************

Intermediate solutions:

"DTM": Dilute three mixed dNTPs to 100μM each in "TM" (we usually 　　　　　　　　　 only use " DTM -C^-")

"LS": Mix 1M HEPES, appropriate DTM and OL in the ratio 25:25:7. 　　　　　　　　　　 Divide into 13μl aliquots and store at -20 C

All solutions can be stored at -20 C.

****************************

Procedure:

Denature DNA fragment to be labelled by incubating in a boiling water bath for 2 minutes. Typically, 50ng DNA is used. This is usually a fragment prepared by "freeze-squeeze" from an agarose gel. Adjust the volume of the DNA fragment to 8.1ul.

Snap-cool in an ice-water bath.

Add 11.4μl "LS", 5μl 10μCi/μl dNTP, 0.5μl 5units/μl DNA polymerase (Klenow fragment) mix and incubate at r.t. for 4 h.

Recover labelled probe by phenol-chloroform extraction and spermine precipitation: Add 75μl H₂O and extract with 100μl phenol-chloroform. Recover the upper aqueous phase to a fresh tube, and re-extract the organic phase with a further 50μl H₂O. Bulk the aqueous phases.

To the aqueous phases, add 100μl 1mg/ml denatured calf-thymus DNA as "carrier" and 14.5μl 100mM spermine. Mix and incubate on ice for 15 min.

Centrifuge in microfuge for 10 minutes and carefully remove and discard the supernatant. This precipitates the DNA but not the unincorporated dNPTs. However, in order to redissolve the DNA it is necessary to exchange out the spermine from the pellet.

Resuspend the pellet in 180μl 75%EtOH-0.3MNaAc-10mMMgAc by vortexing, and incubate on ice for 1 hour.

Spin in microfuge for 10 minutes and carefully remove the supernatant: check that the bulk of the ³²P remains in the pellet with a beta-monitor. Now resuspend the pellet in 180μl 80% EtOH by vortexing, and recover the DNA by spinning in the microfuge for a further 5 minutes.
Carefully remove the supernatant as before, again checking that the labelled DNA remains in the tube. Dry the pellet and dissolve it in 100μl ddH₂O.

Denature the labelled DNA by incubating in a boiling water bath for 2-5 minutes. Snap-cool on ice and add to the hybridisation solution.

Simple method for PCR fragment cleanup

(From Rosenthal A, Coutelle O, Craxton M (1993) Large-scale production of DNA sequencing templates by microtitre format PCR Nucleic Acids Res. 21: 173-4)

Solution (final concentrations):

PEG8000 　　　 26.2%

MgCl₂　　　　　　　 6.6mM

NaOAc 　　　　　 0.6M pH5.2

Mix PCR reaction with an equal volume of the PEG solution and incubate at r.t. for 5 minutes.

Spin at 13,000 rpm for 5 min in a microfuge

Carefully remove the supernatant and wash the (invisible) pellet with 80% EtOH

Spin at 13,000 rpm for 5 minutes, decant the s/n and air-dry.

Dissolve pellet in sterile dd-water or TE

This method precipitates DNA molecules >100-150bp, whilst leaving primers and dNTPs in solution.

(Note: Original citation Waterston R., Martin C., Craxton M., Huynh C., Coulson A., Hillier L., Durbin R., Green P., Shownkeen R., Halloran N., Metzstein M., Hawkins T., Wilson R., Berks M., Du Z., Thomas K., Thierry-Mieg J. & Sulston J. (1992) A survey of expressed genes in Caenorhabditis elegans. Nature Genetics 1: 114-123 uses PEG at a final concentration (after mixing with PCR mix) of 7% and MgCl₂ at 1.75mM).