Archives: Methods

Method:

1. Grow E. coli overnight in 500 mls media that selects for the presence of plasmid.
2. Spin down cells at 8000 rpms for 5 minutes in G3 rotor. Drain thoroughly.
3. Resuspend pellet in 3 mls of ice-cold 25% sucrose in 0.25 M Tris, pH 7.0. Transfer to a 10 ml ultracentifuge tube.
4. Add 0.6 mls of freshly prepared 30 mg/ml lysozyme in H₂0. Mix. Place on ice for 5 minutes.
5. Add 1.2 mls of ice-cold 0.25 M EDTA, pH 8.0. Mix. Place on ice for 5 minutes.
6. Add 1.5 mls of 5 M NaCl. Mix. Immediately add 0.6 mls of 10% SDS. Mix. Leave on ice for 3-4 hours.
7. Pellet lysed cells at 40,000 rpm in a 50Ti rotor at 4^oC in the ultracentrifuge for 45 minutes.
8. Pour supernatent into a 12 ml falcon tube, leaving behind the last bit of viscous clear supernatent (mainly chromosomal DNA). Bring volume to 7.50 mls with sterile water.
9. Add exactly 6.5 grams of CsCl. Mix (wrist action) until dissolved.
10. Add 1 ml of 10 mg/ml Ethidium Bromide (EtBr). Mix. (Caution: EtBr is a mutagen. Wear gloves. Use care. 30 µg = 1 cigarette).
11. Transfer solution to a Beckman quickseal tube (50Ti). Using a 16 gauge needle and a syringe simplifies this. Overlay with parrafin oil. Seal.
12. Spin 48-60 hours at 44,000 RPM in a 50 Ti rotor or 70.1 Ti.
13. Collect the plasmid (lower) band after removing the chromosomal (upper) band. Setup the tube in a clamp over a collection bucket with lots of K-dry around. Wear gloves and a lab coat (EtBr). Visualize with LONG-WAVE UV lamp. Wear a face shield. Collection is done by first puncturing the very top of the tube to allow air in. Then puncuture the tube (3cc syringe with 18 gauge needle) right above the top band and lower the tip into band and suck it off. After all the very viscous portion of the chromosomal band has been removed sick the needle in the opposite wall and get a new 3cc syringe and 18G needle. Stick it in above the plasmid band and suck it off. This is the stuff you want for step 14. Cleanup. (see Maratz (New) p. E8)
14. Extract the EtBr from the plasmid by the addition of an equal part of NaCl-saturated isopropanal (mix equal parts of 5 M NaCl and isopropanol, let sit until the salt settles to the bottom of the tube and there are two phases, use the top phase). Mix. let sit and remove top phase. Repeat until the top phase becomes clear. Do it one more time.
15. Transfer to sterile corex tube that will hold ten times the volume you’ve got. Add two parts water. Mix. Add two parts (new volume, 6 parts original volume) of -20^o C 100% ethanol. Mix. Leave overnight at -20^oC (no longer).
16. Spin down in SS34 (with adaptors) at 8,000RPM for 20 minutes at -20^oC. Drain pellet. Dry. Resuspend in 400 µl of TE or water. Transfer to a 1.5 ml microfuge tube and phenol extract once. Add 60 µl of 3 M Sodium acetate. Mix. Add 800 µl of cold Ethanol. Mix. Leave in -20^oC for a few hours or on dry ice for 10 minutes. Spin down in microfuge in cold room for 15 minutes. Pour off supernatent. Wash with -20^oC 70% Ethanol and spin for 5 minutes. Pour off supernatent and dry pellet in vacuum. Release vacuum slowly (through a paper towel). resuspend in 100 µl TE and take the A₂₆₀, A₂₈₀ and A₃₁₀ of a 1/200 dilution of your prep. 1 A₂₆₀ unit = 50 µg/ml. A₂₆₀/A₂₈₀ ratio should be between 1.5 and 2.0. A₃₁₀ should be very near zero. Alternatively, quantitate with Hoefer TRO₁₀₀ Fluorometer H33228^P dye. (See manufacturer’s method in this manual)

Materials:

Lysis buffer:

Does not need to be sterile but make fresh.

• 50mM Tris-HCl, pH 8.0
• 150mM NaCl
• 5mM MgCl₂
• 3% BSA
• 1ug/ml DNase
• 40ug/ml lysozyme

PBS:

• 140mM NaCl
• 1mM K₂H₂PO₄
• 20mM Na₂HPO₄
• 2.5mM KCl, pH 7.4

Developing Solutions:

See Step 11 to see which ones you need to use.

I. Towbin’s saline:

10mM Tris-HCl, pH 7.4

0.9% NaCl

Substrate solution:

10mg Naphthol AS MX phosphate

10ml 50mM Tris-HCl, pH 8.6

20mg Fast Red

II. Substrate solution:

100 ul of 5mg/ml BCIP

100 ul of 10mg/ml NBT (yellow)

400 ul of 1M MgCl2

9.4 ml of 0.1M Tris pH 9.6

Method:

1. Patch the colonies to be tested on appropriate selective medium.
2. Place orientation marks on both the plate and nitrocellulose and lay a dry filter on top of the colonies.
3. With a gloved finger, gently rub the back of the filter to ensure even transfer of the colonies to the nitrocellulose.
4. Suspend the filter in a glass chamber equilabrated with chloroform for 20 minutes at room temperature.
5. Put the filter in an empty petri dish, add 10ml of lysis buffer and incubate at 37^oC for 45 minutes with shaking.
6. Wash the filter with PBS, squirting the colonies to remove all bacterial debris. Wash twice in PBS for 5 minutes.
7. Add 10ml of primary antibody solution in PBS-1% BSA and incubate at 37^oC for 2 hours with shaking.
8. Wash the filter three times in PBS for 5-10 minutes.
9. Add 10ml of 2nd antibody solution in PBS-1% BSA and incubate as in step 7.
10. Wash the filter with PBS as in step 8.
11. Develop blot using one of the following three methods.

Method I

1. Add 10ml Towbin’s saline and incubate for 10 minutes at room temperature.
2. Add 10ml of Napthol/Fast red substrate solution. (Be careful as the substrate is carcinogenic.) Incubate at room temperature for up to 30 minutes. When the color has reached maximum intensity, wash the filters with distilled water.
3. Dry the filters between paper towel and store away from light.

Method II

1. Add 10 ml of 0.1M Tris pH 9.6 and incubate for 10 min. at room temperature.
2. Add 10 ml of BCIP/NBT substrate solution and incubate at room temperature until color shows up. Wash filters with distilled water.

Method III – For Horseradish Peroxidase Conjugated Antibodies

1. Mix 10 ml of 50 mM Tris pH 7.6 and 0.1 ml chlornapthol stock (prepared by dissolving 0.3 gm of chlornapthol in 10 ml of absolute ethanol. Store -20^oC., stable for at least 1 year)
2. Filter this solution through Whatman No. 1 filter to remove white precipitate.
3. Add 10 ul of 30% H₂O₂ (stored at 4^oC).
4. Positives should develop within 30 minutes. The reaction can be stopped by dumping out developing solution and rinsing with PBS.

**Note: Sodium azide is an inhibitor for HRP.

This is by far the best method for use with ³²P – works every time and the pencil marks develop on the film for easy orientation of the colony lifts!

1. Grow colonies overnight to 1-3 mm in diameter.
2. Blot onto Whatman 541 filter paper. Label top side on front with a pencil. Air dry.
3. In a large glass tray saturate several sheets of Whatman 3MM paper with 0.5 N NaOH. Lay filters colony side up on the saturated paper and leave for 5 minutes. Colonies will turn slimy as they lyse.
4. Transfer filters to a tray with 1 M Tris pH 7.4 and leave for 5 minutes with occasional agitation.
5. Transfer the filters to a tray with 2X SSC and leave for 5 minutes with occasional agitation.
6. Transfer filters to a tray with 95% ethanol. Agitate for 5 minutes. Ethanol will become cloudy. Replace ethanol if doing many filters.
7. Air dry. Do not bake.
8. Place filters in a plastic box with 500 ml 5X SSC and 0.2% SDS. Incubate with shaking at 65C for 30 minutes to remove cell debris.
9. Drain filters and place in a seal-a-meal bag. Add hybridization buffer.
   

   0.25 M NaHPO4	6.25 ml 1 N NaHPO4
   1 mM EDTA	0.05 ml 0.5 N EDTA
   formamide (50%)	12.5 ml
   7% SDS	1.75 g
   water	6.2 ml

10. Prehybridize for 5-10 minutes at 37C. Add denatured probe and hybridize 24 hours at 37C.
11. Wash filters for 5 minutes at RT with 2X SSC.
12. Wash filters for 2 hours at 37C with 250 ml:
    

    0.25 M NaHPO4	75 ml 1 N NaHPO4
    1 mM EDTA	0.05 ml 0.5 N EDTA
    2% SDS	50 ml 10% SDS
    water	124.5 ml

13. Wash for 2 hours with:
    

    0.15 M NaHPO4	37.5 ml 1 N NaHPO4
    1 mM EDTA	0.05 ml 0.5 N EDTA
    1% SDS	25 ml 10%SDS
    Water	187 ml

14. Drain and air dry. Tape filters to a supporting sheet, cover with Saran wrap and autoradiograph.

1. Grow cells overnight in 5ml LB broth (10g tryptone, 5g yeast extract, 5g NaCl per liter).
2. Inoculate into fresh LB broth (50ml will yield 1ml of competent cells; 0.1 ml/transformation) at a 1:200 dilution (OD₅₅₀ should be 0.05 or less). Grow at 37^oC with vigorous shaking in a large flask to facilitate aeration.
3. When cells reach an OD₅₅₀ of 0.2 to 0.4 put them on ice for 10 minutes.
4. Centrifuge the cells 10,000 rpm and 4^oC for 5 minutes in a cold, sterile tube in the SS34 rotor. Discard the supernatant and resuspend the cells in a half volume of sterile, ice-cold (4^oC) 0.1 M CaCl₂. Leave the cells on ice for 20 to 40 minutes.
5. Centrifuge the cells as above. Gently resuspend the cells in 1/50 of the original culture volume in cold, sterile 0.1 M CaCl₂. Leave the cells on ice for 1 to 40 hours (24 hours is optimal for many common strains but spells death to really sick ones). The cells are now competent for transformation.
6. Aliquot 0.1ml of cells to a prechilled eppendorf tube and add the DNA in 10µl or less volume. Make sure to mix the tube gently before aliquoting the cells as they tend to settle at the bottom of the tube. Leave the cells on ice for 20-40 minutes to allow the DNA to adhere to the cells.
7. Heat shock the cells in a 42^oC water bath for 45 to 60 seconds and return to ice for 1 minute.
8. Add 1.2ml of LB broth to the tube, mix and place the eppendorf tube in a small glass test tube and secure with parafilm. Put the tube in the tube roller in the 37^oC room for 1 hour.
9. Plate 0.1ml of the transformed cells on selective plates, spin the remaining cells in the microfuge for 20 seconds and pour off all the media except the last drop (i.e. do not shake out the tube). Resuspend the pellet in the last drop and plate this “neat” on a selective plate. If you anticipate lots of transformants you could plate a 1/10 dilution of the original tube. Incubate the cells overnight at 37^oC. Screen, etc.

Method:

1. Make overnight cultures of the following strains using appropriate antibiotics: E. coli C441 containing helper plasmid: Bacteria with the plasmid to be transferred, 30^oC. and P. aeruginosa recipient strain, 42^oC (ensure 42^oC water bath, as this temperature is required to shut off R/M system).
2. Place 0.1 ml of B in 2 ml of LB and add 0.1 ml of A.
3. Filter the mixture using a 0.45 um Nalgene filter unit. Remove the filter with sterile forceps and place filter side up on an LB agar. Incubate overnight at 30^oC.
4. Resuspend the bacteria in 2 ml of sterile saline (.9% NaCl). Make a 1:100 dilution of this suspension in saline. Spread 100 ul of the undiluted & diluted samples on selection plates. Grow up for 1-2 days at 37^oC.
5. Restreak colonies on selection plates to ensure purity and the presence of the antibiotic marker.

Reference:

O’Toole et al, 1994, Mol. Microbiol. 14: 691-703.

Method:

1. Plate-grown H. pylori are resuspended in BHI to an OD₆₀₀ of 0.6.
2. 1 µg of DNA is added and incubated for 3 h at 37ºC (in a CO₂ incubator).
3. The cells are then spun down and resuspended in 100ul of BHI.
4. The entire 100 µl is spread onto a chocolate blood agar (CBA) plate (no antibiotic) and incubated for 24 h.
5. After 24 h the cells are scraped or washed from the plate and replated on CBA with antibiotic. Single colonies are restreaked for further analysis.
6. PCR can then be used to determine if the gene replacements have occurred as predicted and to confirm that no other rearrangements have occurred.

Method from Poole and Hancock. 1986. Molec Gen Genet. 202: 403-409.

Materials:

• Strain with R68 ts::Tn501 (H526)
• LB (high salt):1% tryptone, 0.5% yeast extract, 0.5% NaCl
• 15 µg/ml HgCl₂
• Carbenicillin 500 µg/ml / tetracycline 100 µg/ml.

Method:

1. Grow strain in LB broth + antibiotics (Carbenicillin and Tetracycline) overnight at 30ºC to select for R68ts::Tn501 plasmid.
2. Do serial dilutions of 10^-1, ^-2, ^-3, ^-4 of the overnight culture.
3. Plate 0.05 ml or 0.1 ml of 10^-3 and ^-4 dilutions onto LB plates (2% agar needed for replica plating) that contain 15 µg/ml HgCl₂. Incubate at 42ºC overnight to select against R68ts::Tn501 plasmid. The HgCl₂ selects for the jumping of Tn501 to the chromosome.
4. Using sterile toothpicks, pick mutants onto fresh LB + HgCl₂ plates. Incubate at 42ºC. overnight.
5. Replicate plates with sterile velvet. Test mutants by colony blot lysis.

1. Photograph ethidium bromide stained gel with a ruler.
2. Soak gel in 0.25 M HCl for 10 minutes (21.5 ml concn HCl/l or 4.3 ml/200 ml)
3. Soak gel in 1.5 M NaCl and 0.5 M NaOH for 30 minutes.
4. Neutralize in 1 M Tris pH 8 and 1.5 M NaCl for 30 minutes.
5. Set up transfer dish using 10X SSC.
6. Invert gel and place onto 3MM paper.
7. Mark nylon membrane with a pencil for later orientation and place onto gel; make sure there are no air bubbles between the gel and the nylon membrane.
8. Put 2 layers of wet 3MM on top of the nylon membrane.
9. Put 2 layers of dry 3MM on top of the wet 3MM paper.
10. Let blot 18-24 hours for chromosomal DNA; 6-8 hours for other DNA (plasmid, etc.).
11. Rinse blot in 2X SSC and UV 30-60 seconds on each side and bake at 80ºC for 2 hours.

NOTE:

If using positively charged nylon, following treatment with HCl, transfer with 0.4 M NaOH and forget about all the other stuff (i.e. use positively charged nylon if at all possible and it will save you a lot of time).

Reference:

Sekar, V. (1987). Biotechniques 5:11-13.

Objective:

This is a quick method to determine the presence of cloned fragments in plasmids after ligation/transformation.

Materials:

Protoplasting buffer:

• 30mM Tris-HCl, pH8.0 (0.33 ml of 1.0 M)
• 5mM EDTA (0.1 ml of 0.5 M)
• 50mM NaCl (0.1 ml of 5.0 M)
• 20% sucrose (5 ml of 40%)
• 50 µg/ml RNAse 1 (50 µl of 10 mg/ml)
• 50 µg/ml lysozyme (50 µl of 10 mg/ml)

Lysis buffer:

• 89mM Tris-HCl, pH8.0 (2 ml of 5X TBE)
• 89mM boric acid
• 2.5mM EDTA
• 2% SDS (2 ml of 10%)
• 5% sucrose (1.25 ml of 40%)
• 0.04% bromphenol blue (4 mg)
• 4.75 ml distilled water

Both solutions may be kept in aliquots at -20^oC.

Method:

1. Aliquot 10 µl of protoplasting buffer into Eppendorf tubes. Do this after pouring an agarose gel (0.6-0.7% agarose in TBE or TAE with SDS is NOT NEEDED!!!).
2. Pick a colony with a toothpick and put the toothpick in the tube and vortex. At this point you can touch a new plate with the same toothpick to obtain a patch colony of the transformant.
3. Preload the gel slots with 4 µl of lysis buffer.
4. Load the protoplast suspension under the lysis buffer. NB: Make sure not to leave the cells in protoplasting buffer longer than 30-40 minutes.
5. Electrophorese in TBE at 20V (minigel) or 40V (large gel) for 15 minutes to allow the cells to lyse completely, then increase the voltage and run until the blue dye migrates to the bottom of the gel.

Stain the gel with 0.5 µg/ml ethidium bromide (if it’s not already included in the gel) for 15 minutes to visualize the bands. Destaining in dH₂O can enhance the photograph of the gel.

1. Run gel as normal in 1X TBE.
2. Visualize band under LONG wave UV.
3. Cut band out with razor blade.
4. Place excised agarose slice in an Eppendorf tube, which has a hole in its bottom (by inserting a hot needle), and which has a small amount of siliconized glass wool covering the opening.
5. Place this tube inside another Eppendorf tube.
6. Spin tubes either at 1/2 maximum speed for 15 minutes or, if not possible, full speed for 5 minutes.
7. Phenol/chloroform the resulting liquid.
8. Repeat with 1/10 volume 3 M NaAc + 2x volume EtOH.