Preparation of Stock PCR Primers

Always take great care not to contaminate the original primer stock - use filter tips and PCR-quality reagents

1. When you receive the lyophilized primer, before opening it be sure to spin it down to insure that the primer pellet is at the bottom of the tube.

2. Prepare the 100 uM primer stock solution in the tube containing the pellet. This tube will be used to make working primer solutions as needed and will be stored at -20°C or below.

To prepare stock:

Determine how many nmoles of primer is in the tube (usually written on the tube or on information accompanying primer order).

Resuspend the pellet in CLEAN 10mM Tris, pH 8.8 buffer to generate the 100uM stock solution. An easy way to do this is to multiply the nmole value by 10 and add that volume in ul to the pellet.

Example: If primer pellet is 8.3 nm, you will add 83 ul 10mM Tris, pH 8.8 buffer to the pellet to resuspend it. The final concentration of primer is 100 uM.

3. To generate the 20 uM working dilution of the primer, dilute the stock solution 1:5 with 10mM Tris, pH 8.8. Example: 10ul stock + 40ul 10mM Tris buffer.

GRAM STAINING - MERCK

Gram staining is a four part procedure which uses certain dyes to make a bacterial cell stand out against its background. The specimen should be mounted and fixed on a slide before you proceed and stain it. The reagents you will need to successfully perform this operation are:

• Crystal violet ( the primary stain)
• Iodine solution (the mordant)
• Decolorizer (ethanol is a good choice)
• Safranin (the counterstain)
• Water (preferably in a squirt bottle)

Before starting, make sure that all reagents, as well as the squirt bottle of water, are accessible. Do this near a sink and wear a lab coat.

1. Place your slide on a slide holder or a rack. Flood (cover completely) the entire slide with crystal violet. Let the crystal violet stand for about 60 seconds. When the time has elapsed, wash your slide for 5 seconds with water. The specimen should appear blue-violet when observed with a naked eye.
2. Now, flood your slide with iodine solution. Let it stand for about a minute as well. Afterwards, rinse the slide with water for 5 seconds and immediately proceed to step 3. at this point, the specimen should still be blue-violet.
3. This step involves addition of the decolorizer, ethanol. This step is somewhat objective because using too much decolorizer could result in a false Gram (-) result. Likewise, not using enough decolorizer may yield a false Gram (+) result. To be safe, add the ethanol dropwise until the blue-violet color is no longer emitted from your specimen. As in the previous steps, rinse the slides with water for 5 seconds.
4. The final step involves applying the counterstain, safranin. Flood the slide with the dye as you did in steps 1 and 2. Let this stand for about a minute to allow the bacteria to incorporate the safranin. Gam (+) cells will incorporate little or no counterstain and will remain blue-violet in appearance. Gram (-) bacteria, however, take on a pink color and are easily distinguishable from the Gram (+) ones. Again, rinse with water for 5 seconds to remove any excess dye.

After completing steps 1—4, blot the slides gently with bibulous paper or allow it to air dry before viewing it under the microscope. DO NOT RUB THE SMEAR!

Decontamination of Dilute Solution of Ethidium Bromide

e.g. electroforesis buffer containing 0.5 ug/ml of ethidium bromide (EtBr)

Method 1 (Lunn and Sansone 1987)

1. Add 2.9 g of amberlite XAD-16 for each 100 mL of solution. Amberlite XAD-16, a nonionic, polymeric absorbent, is available from Rohm and Haas.
2. Store the solution for 12 hours at room temperature, shaking it intermitently.
3. Filter the solution through Whatman No.1 filter, and discard the filtrate.
4. Seal the filter and Amberlite resin in a plastic bag, and dispose of the bag in the hazardous waste.

Method 2 (Bensaude 1988)

1. Add 100 mg of powdered activated charcoal for each 100 mL of solution.
2. Store the solution for 1 hour at room temperature, shaking it intermitently.
3. Filter the solution through a whatman No.1 filter and discard the filtrate.
4. Seal the filter and activated charcoal in a plastic bag and dispose of the bag in the hazardous waste.

Notes:

1. Treatment of dilute solutions of EtBr with Hypochlorite (bleach) is not recommended as a method of decontamination. Such treatment reduces the mutagenic activity af EtBr in the Salmonella/microsome assay by about 1000-fold, but it converts the dye into a compound that is mutagenic in the absence of microsomes (Quillardet and Hofnung 1988)
2. EtBr decomposed at 262 Celcius and is unlikely to be hazardous after incineration under standar condition.
3. Slurries of amberlite XAD-16 or activated charcoal can be used to decontaminate surfaces that become contaminated by EtBr.

Decontamination of Ethidium Bromide Solution (Solutions containing >0.5mg/ml)

Ethidium Bromide (EtBr) is a powerfull mutagen and is moderatelly toxic. Gloves should be worn when working with solutions that contains this dye. After use, these solution should be decontaminated by one of the methods described below.

Decontamination of Concentrated Solutions of EtBr (Solutions containing >0.5 mg/ml)
Method 1
This method (Lunn and Sansone 1987) reduces the mutagenetic activity of EtBr in the Salmonella/ microsome assay by approximately 200-fold.

1. Add Sufficient water to reduce the concentration of EtBr to <0.5>
2. To the resulting solution, add 0.2 volume of fresh 5% hypophosphorous acid and 0.12 volume of fresh 0.5M sodium nitrite. Mix carefully. Check that the pH of the solution is <3.0.
   
3. After incubation for 24 hours at room temperature, add a large excess of 1M sodium bicarbonate. The solution may now be discarded.

note: Hypophosphorous acid is usually supplied as 50% solution, which is corrosive and should be handled with care. It should be freshly diluted immediatelly before use.
Sodium nitrite solution (0.5M) should be freshly prepared by dissloving 34.5 g of sodium nitrite in water to a final volume of 500 ml.

Method 2
This method (Quillardet and Hofnung 1988) reduces the mutagenenic activity of EtBr in Salmonella/microsome assay by approximately 300-fold. However, there are reports (Lunn and Sansone, 1987) of mutagenic activity in occasional batches of "blanks" treated with decontamination solutions.

1. Add sufficient water to reduce the concentration of ETBr to<0.5>
2. Add 1 volume of 0.5 M KMnO4. Mix carefully, and then add 1 volume of 2.5 N HCl. Mix carefully, and allow the solution to stand at room temperature for several hours. Caution:KMnO4 is irritant and is explosive. Solutions containing KMnO4 should be handled in a chemical hood.
3. Add 1 volume 0f 2.5 N NaOH. Mix carefully, and then discard the solution.
   

DNA Electrophoresis on Agarose Gels

Making agarose gel stock (100 mL):

1. Weigh agarose powder on a balance. The amount of agarose needed depends on the concentration of the gel that will be used. The higher the concentration, the resolving ability is also higher.
2. For agarose gel 1% = 1 gram of agarose powder

1.5% = 1.5 grams of agarose powder

2% = 2 grams of agarose powder

3. Mix agarose powder with 1X TAE buffer in a heat-proof bottle. Dissolve it by heating in microwave oven on medium high for several minutes (use no lid or loosen lid while heating).
4. Occasionally during heating, shake the bottle to help the agarose dissolve.
5. Make sure the agarose dissolve completely (the mixture will be absolutely clear and show no traces of undissolved powder). Be careful not to overheat as the agarose mixture will overflow.
6. The agarose mixture is ready to be used directly or stored.

Electrophoresis preparation:

1. Prepare the comb and plate for pouring melted agarose. Use an appropriate comb depending on the number of samples that will be run.
2. Wait until the liquid agarose mixture cools down to around 60 ^oC (cool enough to not scorch your hand but not too cool that it partly solidifies). DO NOT POUR BOILING AGAROSE MIXTURE ON PLATE!
3. Pour agarose mixture on plate; the volume will depend on the volume of samples to be put in wells. For a reference, 30 mL of agarose mixture poured on plate with a standard 8-well comb will hold a maximum of 20µL of sample in each well.
4. Leave the agarose gel to set (around 10—15 minutes).
5. When the gel is set, take out the comb and put the plate in electrophoresis tank.
6. Pour enough 1X TAE buffer to cover the gel and fill the tank (approximately 250 mL buffer).

Loading samples and running electrophoresis:

1. Take out the samples, loading dye (2X), and DNA marker/ladder from fridge/freezer. Thaw completely.

2. Mix samples with loading dye on a parafilm sheet. As a guide, 2 µL of loading dye is enough to hold down 4—5 µL of sample. For larger amounts of samples, increase the amount of loading dye accordingly. Pipette up and down using a micropipette to mix the sample and loading dye completely.

3. Carefully pipette the sample/dye mixture into each well. After all samples are loaded into wells, load 4 µL of DNA ladder into one well. Note: the DNA ladder being used here is ready-to-use (already mixed with loading dye). If not, mix it with appropriate amount of loading dye first before pipetting it into the well.

4. Assemble the lid on tank and connect it to a power source. Make sure the side of gel that has wells is placed on the side of tank that is connected to the cathode (negative) of power source, as DNA is negatively charged and will travel to positively charged side (anode).

5. Set the current to maximum and set the voltage to 60-80 volts.

6. Run the electrophoresis for around 1—2 hours or until the loading dye reaches ¾ of gel (higher concentration of gel and lower voltage means longer running time). Do not let it run for too long or the DNA will be lost in the buffer.

Visualising DNA fragments:

1. When electrophoresis is finished, turn off power and open the tank. Take out the gel and put it on a container.
2. Pour EtBr (ethidium bromide) solution (30 µL in 500 mL of water) over the gel until it is completely soaked.
3. Soak the gel for 10—15 minutes. If the ethidium bromide solution is old or has been re-used many times, leave the gel soaked for longer.
4. Afterwards, pour back EtBr solution back in its bottle and wash the gel with tap water for 2—3 minutes.
5. Visualise DNA by viewing the gel under UV light.
6. If the DNA fragment is too faint, soak the gel again in EtBr solution. If the gel is too bright (soaked too long in EtBr), increase washing time to 5—6 minutes.
7. Photograph the gel with a manual polaroid camera or with GelDoc System.

Competent Cells

CaCl₂ METHOD

(All activities are done in aseptically in laminar flow cabinet, except for centrifuging).

1. Prepare overnight culture – pick a colony of E. coli on agar plate and grow in 3 mL LB broth. Incubate at 37 ^oC overnight.
2. Use 1 mL overnight culture to inoculate 100 mL LB broth in 500 mL erlenmeyer flask.
3. Incubate at 37 ^oC with shaking (200 rpm).
4. Grow to OD 600 = 0.4—0.5 (optimal for DH5α). Usually takes about 3 hours.
5. Make 2 x 50 mL aliquots in oakridge tubes, centrifuge at 500 rpm for 15 minutes. Discard supernatant.
6. Resuspend pellet in 10 mL ice cold CaCl₂ solution.
7. Centrifuge at 5000 rpm for 15 minutes, discard supernatant.
8. Resuspend each pellet in 1 mL ice cold CaCl₂ solution.
9. Combine and make 200 µL aliquots in sterile 1.5 mL microcentrifuge tubes.
10. Snap freeze on dry ice/ETOH.
11. Store at -70 ^oC, labelled CC.

For transformation (heat-shock method):

1. Mix 100—200 µL of CC with 1 µL plasmid DNA (or 10 µL ligation mixture) in a sterile 1.5 mL microcentrifuge tube. Incubate on ice (or put in freezer) for 30 minutes.
2. Heat shock treatment: put tube in dry bath or water bath at 42 ^oC for 2 minutes.
3. Immediately put on ice for 10 minutes.
4. Cell/DNA mixture is spread on agar plate (plus ampicillin or other antibiotic). Incubate at 37 ^oC overnight.

TSS METHOD (1)

1. Inoculate 3 mL of LB broth with an E. coli colony from agar plate. Incubate at 37 ^oC overnight, with shaking.
2. Dilute 1:100 fresh overnight culture of bacteria into pre warmed LB broth and incubate cells with shaking (225 rpm) to an OD 600 of 0.3—0.4 (approximately 3 hours).
3. Add an equal volume of ice-cold 2X TSS and mix gently. [TSS is LB broth with 10% PEG (MW 3350-8000), 5% DMSO, and 20-50 mM Mg ²⁺ (MgSO₄ or MgCl₂) at a final pH of 6.5].
4. For long term storage, cells are frozen immediately in a dry ice/ethanol bath and stored at -70 ^oC.

TSS METHOD (2)

1. Inoculate 50 mL LB broth with 0.5 mL fresh overnight culture and incubate at 37 ^oC with shaking until OD 600 reaches 0.3-0.4.
2. Spin to pellet cells after mixing (5000 rpm, 3 minutes) and add 2 mL of 1 X TSS.

For transformation (1):

1. A 0.1 mL aliquot of competent cells pipetted into a cold polypropylene tube containing 1 µL (100 pg) of plasmid DNA or ligation mixture, and cell/DNA suspension is mixed gently. [When frozen cells are used, cells are thawed slowly on ice and used immediately].
2. The cell/DNA mixture is incubated for 5—60 minutes at 4 ^oC.
3. A 0.9 mL aliquot of TSS (or LB broth) plus 20 mM glucose is added, and cells are incubated at 37 ^oC with shaking (225 rpm) for 1 hour to allow expression of the antibiotic resistance gene.
4. Transformants are selected with standard methods.

another Molecular Biology Reagents

PMSF 100mM

Dissolve 17.4 mg/mL in isopropanol

Store at -20C

30% Acrylamide/bis

72.5 mL Acrylamide 40%

1 g bis-acrylamide

Dissolve in water until 100 mL

TBS

10 mM Tris-HCl pH 7.5

15 mM NaCl

Sterilize by autoclaving

IPTG 100 mM

For 20 mL, dissolve 476.6 mg IPTG in 20 mL water

TTBS

TBS with 0.1% of Triton X-100

1 Kb dna ladder marker

Stock = 500 ug/uL

Dilute to 125 ng/uL

For work : 100 uL marker + 100 uL 1XTE + 200 uL 2XLB

100 bp dna ladder marker

Stock = 500 ug/uL

Dilute to 125 ng/uL

For work : 100 uL marker + 100 uL 1XTE + 200 uL 2XLB

1 Kb dna ladder marker

Stock = 250 ug (1 ug/uL)

Dilute to 125 ng/uL (1:8)

For work : 50 uL marker + 150 uL water + 200 uL 2XLB

Tris-Glycine Electroforesis Buffer 5X Stock

15.1 g tris base

94 g Glycine

900 mL water

50 mL SDS 10%

Addjust volume with water until 1000 mL

2XSB

100 mM DTT (Stock 1M)

2% SDS

80 mM Tris-HCl pH 6.8

0.006% w/v bromophenol blue

15% Glycerol

Agarose TOP (100 mL)

LB powder for 100 mL

100 mg MgCl2

700 mg Agarose

Sterilize with autoclaving, store in refrigerator

Inclusion Body wash Solution 1 (Cold)

2 M urea

20 mM Tris

0.5 M NaCl

2% Triton X-100

Adjust to pH 8.00

Inclusion Body Wash Solution 2 (Cold)

20 mM Tris

0.5 M NaCl

2% Triton X-100

Adjust to pH 8.00

Solubilisation Buffer

20 mM Tris

0.5 M NaCl

5 mM Imidazole

6 M Guanidine HCl

1 mM Beta-mercaphtoethanol (BME)

Adjust to pH 8.00

(Concentration of BME might have to be optimized)

Equilibration buffer

300 mL ethanol 95%

95 mL Glycerol 50%

555 mL water

Molecular Biology Reagents

Luria Broth LB

(prepare with dd H2O, sterlilized by autoclaving)

1% w/v bacto-tryptone

0.5% w/v yeast extract

1% w/v NaCl

pH adjusted to 7.0 using NaOH 0,6 N

Solution 1 ( Cell Resuspension Solution)

50 mM Glucose (4.5 mL; stock 20%)

25 mM Tris pH 8 (2.5 mL; stock 1 M)

10 mM EDTA (10 mL; stock 0.1 M)

Solution 2 ( Cell Lysis Solution, Fresh)

2 M NaOH 1 mL

10 % SDS 1 mL

Dd H2O 8 mL

Solution 3 ( Neutralisation Solution)

5 M potassium acetate 60 mL

Glacial cetic acid 11.5 mL

H2O 28.5 mL

2X TSS (Competent cells media, TSS methods)

TSS=LB + 10% PEG + 5% DMSO + 20-50 mM MgCl2

pH=6.5

Glycerol-CaCl2 Solution (competent cells media, CaCl2 methods)

22.5 mL 0.4 M CaCl2

28.1 mL 80% Glycerol

99.4 mL H2O

TAE 50X/L

242 g Tris Base

57.1 mL Glacial acetic acid

100 mL 0.5 M EDTA pH 8

TBE 5X/L

54 g Tris Base

27.5 g Boric Acid

20 mL 0.5 M EDTA pH 8

EtBr

Stock Solution 10 mg/mL ( 1 tablet / 1 mL H2O)

Working Solution 0.5 ug/mL (250 mL H2O)

Double Detergent Lysis Bufer (100 mL)

50 mM Tris-HCl pH 8

150 mM NaCl

0.1% SDS

1% IGEPAL

SDS-PAGE Gel

12% resolving gel (1 gel)

3.4 mL H2O

2.5 mL 1.5M Tris-HCl pH 8.

50 uL SDS 20%

4 mL Acrylamida/bis

50 uL APS 10%

5 uL TEMED

12% stacking gel (2 gel)

3.075 mL H2O

1.25 mL Tris-HCl pH 6.8

25 uL SDS 20%

0.67 mL Acrylamida/bis

25 uL APS 10%

5 uL TEMED

8% Resolving gel (2 gel)

9.3 mL H2O

5.0 mL Tris-HCl pH 8.8

0.1 mL SDS 20%

5.3 mL Acrylamida/bis

0.2 mL APS 10%

0.012 mL TEMED

8% Stacking gel = 12% Stacking gel

Low Toxicity Staining Solution

0.25 g coomassie blue R250

100 mL ethanol

100 mL water

Stir 1 hour

+ 25 mL Acetic Acid

+water until 250 mL

Stored at room temperature (Alumunium foiled bottle)

Destaining Solution

400 mL ethanol

100 mL acetic acid

500 mL water

Stored at room temperature