Isolation of drug resistant mutants using UV – Gradient plate and Replica plating

 

 Isolation of drug resistant mutants using UV – Gradient plate and Replica plating

AIM:   To become familiar with gradient plate method for isolating antibiotic resistant mutants and to perform replica plating procedure for isolating and identifying Auxotrophic mutants.

Principle

 Mutations often inactivate a biosynthetic pathway of the microorganism, and frequently make a microorganism unable to grow on a medium lacking an adequate supply of the pathway’s end product. Based on this principle microorganism are classified as Prototrophic and Auxotrophic. Prototrophic organisms (wild type) have the same nutritional requirements as that of their ancestors. They need only inorganic salts, an organic energy source such as sugar, fat, protein and water to survive and grow. That is, the Prototroph’s need only "Minimal medium" for their growth and survival. Auxotrophic mutants are unable to grow without one or more essential nutrients. Auxotrophs are mutant for particular nutrient synthesis pathway enzymes. Such an error is known as an inborn error of metabolism, whether it occurs in a bacterium or a eukaryote. An auxotroph can be grown only on an enriched medium that provides the particular nutrient that the mutant cannot metabolize on its own.

Gradient Plate Technique: 

            The gradient plate consists of two wedges like layers of media: a bottom layer of plain nutrient agar and top layer of antibiotic with nutrient agar. The antibiotic in the top layer, diffuse into the bottom layer producing a gradient of antibiotic concentration from low to high.  A gradient plate is made by using Streptomycin in the medium. E. coli, which is normally sensitive to Streptomycin, will be spread over the surface of the plate and incubated for 24 to 72 hours. After incubation colonies will appear on both the gradients. The colonies develop in the high concentration are resistant to the action of Streptomycin, and are considered as Streptomycin resistant mutants. For isolation of antibiotic resistant of gram negative enteric bacteria, the antibiotics commonly used are Rifampicin, Streptomycin, and Erythromycin etc.

 

 

 

Replica Plating Method:

The technique was developed by Joshua and Esther Lederberg in 1952 for providing the direct evidence for the existence of pre-existing mutations. This technique isolates both nutritional mutants and antibiotic resistant mutants. Their actual experiment concerned with replicating master plates of sensitive cells to two or more plates containing streptomycin. Replica plating allows the observation of microbes under a series of growth conditions. The bacteria are grown in an environment that is not selective for given mutation.  This technique is used to transfer the members of each colony to a selective environment. A simple velveteen covered colony transfer device is used to transfer the colonies in nutrient agar medium supplemented with or without a particular antibiotic or nutrient.  The fibers of velvet act as fine inoculating needles, picking up the bacterial cells from the surface of this master plate. The velvet with its attached microbes is then touched to the surface of a sterile agar plate, inoculating it. In this manner, microbes can be repeatedly stamped onto media of differing composition. By comparing the presence of colonies following incubation we can indirectly determine the mutant colonies by their absence in the selective environment. A colony that develops on a complete medium fail to develop on a minimal medium that lacks a specific growth factor, the occurrence of a nutritional mutant is indicated. The microbes that do not grow on the minimal medium represent auxotrophic strains. The microbes that do not grow on the minimal medium represent auxotrophic strains.

 

 

 Replica plating technique

 

 

 

Materials required: Gradient Plate Technique

 

1.       24 hour old nutrient broth culture of Escherichia coli.

2.      Two nutrient agar deep tubes (10 ml per tube/culture).

3.      1% Streptomycin sulphate solution (100 µg/ml).

4.      A beaker with 90% ethanol.

5.      Sterile Petri plates.

6.      Sterile 1 ml pipette.

7.      Glass rod spreader.

8.      Water bath.

Procedure: I) Preparation of gradient plate:

 

1.      Melt two nutrient agar plates maintained at 96⁰C and cool to 55⁰ C

2.      Pour the contents of one agar tube into a sterile petriplate. Allow the medium to solidify in a slanting position by placing either a glass rod under one side.

3.      After the agar medium is solidified remove the glass rod and place the plate in the horizontal position.

4.      Pipette out 0.1mL of 1% Streptomycin solution into the second tube of the second nutrient agar medium.

5.      Rotate the tube between the palms and pour contents to cover the gradient layer agar and allow to the medium to solidify on a level table.

6.      Label the low and high antibiotic concentration area on the bottom of the plate.

 

II) Inoculation of culture:

 

1.      Pipette out 200µl (0.2ml) of the overnight Escherichia coli culture onto the gradient plate after 24 hours of its preparation.

2.      Spread the inoculums evenly over the agar surface With a sterile bent glass rod by rotating the plate.

3.      Incubate the inoculated plate in an inverted position at 37^o C for 48-72 hours. 

4.      Observe the plate for appearance of E.coli colonies in the area of low streptomycin concentration (LSC) and high streptomycin concentration (HSC) and record the results.
 

Results: Colonies which appear in the area of high concentration streptomycin region will be streptomycin resistant mutants.

Replica plating method:

 Materials required:

1.      24 hour old nutrient broth culture of Escherichia coli.

2.      Minimal salt agar with glucose.

3.      Three 10ml Nutrient agar deeps.

4.      1% Streptomycin sulphate solution (10mg /100ml of sterile water).

5.      Sterile petridishes.

6.      Sterile velveteen colony carrier.

7.      Glass rod or wooden dowel stick .

8.      Beaker with 95% ethanol.

9.      Bent glass rod.

10.  Quebec colony counter

 

 

Procedure:

DAY 1

1.      Melt the nutrient agar deeps tubes in a hot water bath maintained at 96⁰ C.

2.      Allow the molten medium to cool to 55⁰ C.

3.      Pour the molten agar medium to two sterile petriplates and allow to solidify in a horizontal position.

4.      Add 0.1% of Streptomycin, using sterile pipette into the third tube of molten nutrient agar (maintained at 55⁰C ), properly mix by rotating between the hands and pour the contents into a sterile petriplate. Allow to solidify.

5.      Add 200 µL of(0.2mL) of the the E.coli test culture to the surface of the nutrient agar plate.

6.      Using an alchohol dipped and flamed bent glass rod spread the inoclum evenly on the plate.

7.      Incubate the plate in an inverted position for 24-48 hours at 37 ⁰ C.

8.      After incubation observe the colonies of E.coli on the plate and this plate was considered as the master plate.

DAY 2

 

1.      A reference mark (at 12 O'clock position) was noted on the bottom of the master plate, plate with nutrient agar and plate supplemented with Streptomycin.

2.      The sterile velveteen colony carrier was carefully lowered and gently pressed onto the colonies of the E.coli on the master plate.

3.      Without altering the position of the carrier, the sterile velveteen was gently pressed on to the nutrient agar plate followed by the Nutrient agar plate supplemented with Streptomycin.

4.      Incubate both the inoculated plates, nutrient agar and streptomycin agar plates in an inverted position for 48-72 hours at 37⁰C. Master plate is refrigerated.

5.      Following incubation number of colonies in the replica plates with nutrient agar and streptomycin agar was counted using Quebec Colony Counter.

6.      The colonies appearing on the nutrient agar plates and Streptomycin plates were noted and compared.

Results:

E.coli colonies which appear on the Streptomycin supplemented agar was confirmed as Streptomycin resistant mutants and the number of colonies were counted using Quebec Colony Counter.

 

 

WIDAL TUBE TEST

 

WIDAL TUBE TEST

Introduction

Widal test is a serological test which is used for the diagnosis of enteric fever or typhoid fever. The test was developed by Greembaum and Widal in 1896. Typhoid or enteric fever is caused by a gram negative bacteria Salmonella enterica (Salmonella Typhi or Salmonella Paratyphi), found in the intestine of man. Salmonella paratyphi also causes Typhoid but of a milder form.

Salmonella possess O antigen on their cell wall and h antigen on their flagella. On infection, these antigen stimulates the body to produce specific antibodies which are released in the blood. The Widal test is used to detect these specific antibodies in the serum sample of patients suffering from typhoid using antigen-antibody interactions. These specific antibodies can be detected in the patient’s serum after 6 days of infection (fever).

Salmonella Typhi possesses O antigen on the cell wall and H antigen on flagella. Salmonella Paratyphi A and S. Paratyphi B also possess O antigen on their cell wall and but have AH and BH antigen on their flagella respectively.

Principle

 Widal test is an agglutination test in which specific typhoid fever antibodies are detected by mixing the patient’s serum with killed bacterial suspension of Salmonella carrying specific O, H, AH and BH antigens and observed for clumping ie. Antigen-antibody reaction. The main principle of Widal test is that if homologous antibody is present in patient’s serum, it will react with respective antigen in the suspension and gives visible clumping on the test slide or card.

Requirements

i) Fresh serum, stored at 2-8° Serum should not be heated or inactivated.

ii) The complete kit containing five vials containing stained Salmonella antigen

S. Typhi———-O antigen,

S. Tyhhi———- H antigen

S. Paratyphi —–AH antigen

S. Paratyphi —–BH antigen

iii) Widal positive control

iv) Widal test card or slide

v) Applicator stick

 

Procedure

1.      Bring all reagents to room temperature and mix well.

2.      Prepare 4 sets of test tubes for individual antigen. Each set contains 1- 8 tubes.

3.      Add 1.9 ml of 0.85% sterile saline to tube no. 1 of each antigen set.

4.      To tube no. 2-8 of all sets add 1 ml of physiological saline.

5.      To tube No. 1 of all sets add 0.1 ml of test sample to be tested and mix well.

6.      Transfer 1 ml of the diluted serum sample from tube No. 1 to tube No. 2 and mix well.

7.      Transfer 1 ml of the diluted serum sample from tube No. 2 to tube No. 3 and mix well. Continue this serial dilution till tube No. 7 in each set of antigen.

8.      Discard 1.0 ml of the diluted serum from tube No.7 of each set.

9.      So the dilutions of the serum sample from tube No. 1 to 7 respectively in each antigen set are 1:20, 1:40,1:80, 1:160, 1: 320, 1:640, 1: 1280.

10.  Tube no. 8 is negative control with 0.85% sterile saline.

11.              To one set i.e. from tube no.1- 8 add 50 µl of Salmonella typhi ‘O’ antigen.

12.  In second set i.e. from tube no.1- 8 add 50 µl of Salmonella typhi ‘H’ antigen.

13.  Respectively for third and fourth sets, add Salmonella paratyphi ‘AH’ and Salmonella paratyphi ‘BH’ to all tubes from 1-8.

14.  Mix well, cover and incubate these tubes overnight at 37 degree Celcius (approximately 18 hours).

15.  After incubation dislodge the sediment and observe for agglutination.

Interpretaton :

The antibody titre of the test sample is its highest dilution that gives a visible agglutination. Agglutinin titre greater than 1:80 is considered as significant infection and low titres indicate absence of infection.

 

PLASMID ISOLATION FROM BACTERIA

 

 

PLASMID ISOLATION FROM BACTERIA

 

Introduction

In virtually all bacterial species plasmids exist. These accessory genetic elements typically account for only a small fraction of a bacterial genome corresponding roughly to a range between 1 and 200 kb. Extremely large plasmids with sizes far beyond 200 kb are also known. Plasmids of more than 50 kb might be characterized as “large”, plasmids of less than 10 kb as “small”. The aim of this compilation is to describe some fast methods for plasmid isolation leading to “crude lysates”, the quality of which being sufficient for analytical purposes, mainly agarose gel electrophoresis. By using few microliters of crude lysates for agarose gel electrophoresis, the electrophoretic separation allows conclusions on

* The presence of plasmid DNA,

* The determination of the molecular weight(s),

* The amount of plasmid DNA due to the band intensity and on

* The purity of the crude lysate.

The fast methods described here are often suitable for plasmid screenings from bacteria in E. coli.

 

 Principle

The procedures are based on the fact that plasmids usually occur in the covalently closed circular (supercoiled) ccc configuration within the host cells. After gentle cell lysis all intracellular macromolecules have to be eliminated whereas plasmid DNA is enriched and purified. The smaller a plasmid the easier is the isolation of intact ccc molecules. DNA is very sensitive to mechanical stress, therefore shearing forces caused by mixing/vortexing or fast pipetting must be avoided as soon as cell lysis occurs. All mixing steps during and after cell lysis should be performed carefully by inverting the tubes several times (8-10 fold). Especially in case of larger plasmids it is recommended to cut off the ends of plastic pipette tips to minimize shearing forces. Gloves should be worn in order to prevent contamination with DNases. Autoclaved (DNase-free) buffer solutions, tubes and tips should be used. If phenotypic markers of a plasmid (e.g. antibiotic resistances) are known, it is recommended to grow the cells under selective pressure to avoid plasmid loss. If necessary, small plasmids of Escherichia coli can easily be amplified using chloramphenicol. This results in several thousand plasmid copies per cell leading to high DNA quantities (Clewell, 1972). Large plasmids are maintained with only one copy per host chromosome: visible DNA bands are more difficult to get.

For plasmid isolation, bacterial cultures should be grown to late logarithmic/early stationary phase. It is important to remove the supernatant completely after centrifugation from the cell pellets. Tris buffer is the typical buffering substance for DNA with buffering capacity in the slightly alkaline range in which DNA can also be stored best (pH 7.5-8.2). EDTA is an important substance in plasmid preparations because it inhibits nuclease activity. For long-term storage, plasmid DNA should be frozen in aliquots of storage TE buffer. Repeated thawing and freezing of DNA should be avoided.

Ethanol precipitation of plasmid DNA

Measure the volume of the aqueous DNA solution and mix gently with (10% v/v) 3 M Na-acetate, pH 5.2, then add double of the total volume of pure ethanol (cooled to -20C), mix and leave for 10 min in crushed ice. Spin for at least 30 min at room temperature. DNA precipitation is not enhanced by long or low temperature incubation, whereas an extended centrifugation time results in good DNA recovery.

RNase treatment

Prepare 100 ml of the following sterile TE buffer: 0.01 M Tris, pH 7.5, 0.001 M EDTA. Mix 1 mg of RNase A with 1 ml of this TE buffer in an Eppendorf tube and incubate for 20 min in a boiling water bath to eliminate DNases. Cool to room temperature, add the RNase solution to the remaining 99 ml of the same TE buffer. This RNase buffer can be stored at 4°C for a long time and is a good storage buffer for plasmid DNA. RNase is a very stable enzyme and cleaves RNA within few minutes at room temperature.

Gel electrophoresis

Immediately before loading a gel, mix 8 μl of DNA sample with 2 μl of loading buffer (0.05 M EDTA, 20% Ficoll, 0.25% bromophenol blue, in H20).

When using a horizontal electrophoresis apparatus (horizontal apparatus is the usual and better type of electrophoresis), for quick analytical gels, mini-gels on glass slides can be prepared as follows: about 25 ml of 0.8-1.0% low electroendosmosis (EEO) agarose in TBE buffer (0.089 M Tris, 0.089 M boric acid, 0.0025 M EDTA) are poured on a glass slide of approx. 10 x 7 cm. Depending on the electrophoresis comb used, up to 14 samples can be run. The same TBE buffer is used as electrophoresis buffer. Usually, the electrophoretic separation is done at 30-90 V for 2-6 hours (to be tried out). For visualization of DNA bands and photography, intercalating dyes like ethidium bromide are used: After staining for 30-60 minutes in the dark, DNA bands can be made visible under short wave length UV light.

Procedure:

1. Hot alkaline method for all plasmid sizes and bacteria

                         

Centrifuge 2-3 ml of culture, resuspend pellet in 1 ml of solution containing 0.04 M Tris-acetate, pH 8.0 (adjust pH with glacial acetic acid) and 2 mM EDTA

 

Add 2 ml of lysis buffer (0.05 M Tris, 3% SDS, pH 12.50, adjusted with 2 N NaOH) and mix

 

Incubate at 60-68°C for 30-45 min (strain dependent)

 

Add to hot samples 6 ml of phenol/chloroform (1:1) and mix gently to complete emulsification

 

Separate phases by centrifugation at 10.000 x g for 15-20 min at RT and transfer the upper aqueous phase carefully (avoid interphase which contains debris) to new tube containing 1 volume of chloroform. Mix and centrifuge again for separation of phases

 

Recover aqueous phase and use directly for agarose gel

 

2. Lysozyme method for various Gram-negative bacteria

                         

Centrifuge 10 ml of culture, resuspend pellet in 1.4 ml of the following TE buffer: 0.01 M Tris, pH 8.5 and 1 mM EDTA. Transfer to Eppendorf tubes and spin for 3 min

 

Resuspend pellet in 0.4 ml of solution (15% sucrose, 0.05 M Tris, pH 8.5, 0.05 M EDTA), mix vigorously, and cool on ice

 

Add 0.1 ml of freshly prepared lysozyme (5 mg/ml in TE buffer used above), mix carefully and incubate on ice for 20-40 min

 

Add 0.3 ml of pre cooled Triton buffer (0.1% Triton X-100, 0.05 M Tris, pH

 

8.5, 0.05 M EDTA), incubate on ice for 20 min and centrifuge at 4°C for 4 min

 

Transfer clear supernatant into new tube and add 4 μl of diethyloxydiformiate, mix gently

 

Incubate for 15 min at 70°C, cool for 15 min to RT, then incubate on ice for 15 min

 

Centrifuge for 4 min, transfer supernatant into new tube, fill up with ¬20°C ethanol for DNA precipitation, mix gently

 

Centrifuge for at least 30 min at RT, dry pellet in vacuum dessiccator and resuspend in storage TE buffer or in RNase buffer before use.

 

3. Lysis of cells from single colonies on agarose gel

Transfer 1-2 freshly grown single colonies with a toothpick into 20 μl of cold buffer (0.025 M Tris, pH 8.0, 25% sucrose, 0.250 M EDTA, 7% Ficoll 400)

 

Add 20 μl of freshly prepared lysis solution (0.1 mg/ml of lysozyme, 10 μl/ml of RNase A, in the above buffer), mix well and immediately fill 10-15 of the mixture into the well of an agarose gel which contains 0.5% SDS

 

Add as “upper layer” onto the cell lysate 10 μl of the following solution: 0.025 M Tris, pH8.0, 10% SDS, 25% sucrose, 0.07% bromophenol blue

 

After 15-30 min apply low voltage (half of usual voltage) for 30 min, and then apply usual electrophoretical conditions

 

4. Isolation procedure for all plasmid sizes from all bacteria

Centrifuge 2 ml of a culture and wash pellet in 2 ml of the following TE buffer: 0.05 M Tris, pH 8.0, 0.01 M EDTA. Resuspend in 40 μl of the same TE buffer

 

Fill 0.6 ml of freshly prepared lysis buffer (TE buffer used above with 4%SDS, pH adjusted to 12.45) into Eppendorf tube and add the cell suspension to the lysis buffer, mix gently

 

Complete lysis by incubating at 37°C for 20-30 min

 

Add 30 μl of 2 M Tris, pH 7.0 for neutralization, mix gently

 

Add 024 ml of 5 M NaCl for precipitation of chromosomal DNA and protein and incubate on ice for 4 hrs

 

Centrifuge for 10 min and transfer supernatant into new tube for ethanol precipitation (as usual) or for previous extraction with phenol/chloroform.