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Method for preparation of LB agar plates with Ag+?

Method for preparation of LB agar plates with Ag+?



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I need a method for preparing LB agar plates with 1 mM concentration of AgNO3. I can not seem to find a method online, does anyone know of one that I could use?

Here are my main questions regarding the method:

a. Do I add the silver before or after I autoclave?

b. do I add the silver in a liquid form for powder?

Thanks.


You add the silver after autoclaving. Just make sure that you make your stock concentration of silver in sterile water to avoid unwanted contamination. Do it in liquid form; it is easier to dilute from a stock.


Culture Media for Cultivation of Anaerobic Bacteria: 4 Types

The following points highlight the top four types of culture media used in the cultivation of anaerobic bacteria. The culture medium are: 1. Special Anaerobic Culture Media 2. Anaerobic Chamber 3. Anaerobic Bags or Pouches 4. Anaerobic Jars.

Type # 1. Special Anaerobic Culture Media (Prereduced Media):

Of all the methods available for the cultivation of anaerobic bacteria, exclusion of oxygen from the medium is the simplest method. During preparation, the liquid culture medium is boiled by holding in a boiling water both for 10 minutes to drive off most of the dissolved oxygen.

Liquid media soon become aerobic thus a reducing agent (e.g., cysteine 0.1%, ascorbic acid 0.1%, sodium thioglycollate 0.1%), is added to further lower the oxygen content.

Oxygen-free N2 is bubbled through the medium to maintain anaerobic condition. The medium is then dispensed into tubes, which are stoppered tightly and sterilized by autoclaving. Such tubes can be stored for many months before being used. During inoculation, the tubes are continuously flushed with oxygen free CO2 by means of gas cannula, re-stoppered, and incubated (Fig. 16.18).

Cooked meat broth (CMB original medium known as ‘Robertson’s bullock-heart medium’) has a special place in anaerobic bacteriology, and thioglycollate broth and its modifications are also very useful. CMB is suitable for growing anaerobic bacteria in air and also for the preservation of their stock cultures.

The inoculum of the bacterium is introduced deep in the medium in contact with the meat. Meat particles are placed in 30 ml bottles to a depth of about 2.5 cm and covered with about 15 ml broth.

However, some other media which can be used for recovering anaerobes are Brucella blood agar, Bacteroides bile aesculin agar, phenylethyl alcohol agar, kanamycin blood agar, etc. Anaerobic bacteria have special nutritional requirements for vitamin K, haemin and yeast extract, and all primary isolation media for anaerobes should contain these three ingredients.

Type # 2. Anaerobic Chamber:

Anaerobic chamber is an ideal anaerobic incubation system, which provides oxygen- free environment for inoculating media and incubating cultures. It refers to a plastic anaerobic glove box that contains an atmosphere of H2, CO2, and N2. Glove ports and rubber gloves are used by the operator to perform manipulations within the chamber. There is an air-lock with inner and outer doors.

Culture media are placed within the air-lock with the inner door. Air of the chamber is removed by a vacuum pump connection and replaced with N2 through outer doors.

The culture media are now transferred from air-lock to the main chamber, which contains an atmosphere of H2, CO2, and N2. A circulator fitted in the main chamber circulates the gas atmosphere through pellets of palladium catalyst causing any residual O2 present in the culture media to be used up by reaction with H2.

When the culture media become completely anaerobic they are inoculated with bacterial culture and placed in an incubator fitted within the chamber. The function of CO2 present in the chamber is that it is required by many anaerobic bacteria for their best growth. A schematic representation of an anaerobic chamber showing its various parts is given in Fig. 16.19.

Type # 3. Anaerobic Bags or Pouches:

Anaerobic bags or pouches make convenient containers when only a few samples are to be incubated anaerobically. They are available commercially. Bags or pouches have an oxygen removal system consisting of a catalyst and calcium carbonate to produce an anaerobic, CO2-rich atmosphere.

One or two inoculated plates are placed into the bag and the oxygen removal system is activated and the bag is sealed and incubated. Plates can be examined for growth without removing the plates from bag, thus without exposing the colonies to oxygen.

But as with anaerobic jar, plates must be removed from the bags in order to work with the colonies at the bench. These bags are also useful in transport of biopsy specimen for anaerobic cultures.

Type # 4. Anaerobic Jars (or GasPak Anaerobic System):

When an oxygen-free or anaerobic atmosphere is required for obtaining surface growth of anaerobic bacteria, anaerobic jars are the best suited. The most reliable and widely used anaerobic jar is the Melntosh-Fildes’ anaerobic jar. It is a cylindrical vessel made of glass or metal with a metal lid, which is held firmly in place by a clamp (Fig. 16.20).

The lid possesses two tubes with taps, one acting as gas inlet and the other as the outlet. On its under surface it carries a gauze sachet carrying palladium pellets, which act as a room temperature catalyst for the conversion of hydrogen and oxygen into water. Palladium pellets act as catalyst, as long as the sachet is kept dry.

Inoculated culture plates are placed inside the jar and the lid clamped tight. The outlet tube is connected to a vacuum pump and the air inside is evacuated. The outlet tap is then closed and the gas inlet tube connected to a hydrogen supply. Hydrogen is drawn in rapidly. As soon as this inrush of hydrogen gas has ceased the inlet tube is also closed.

After about 5 minutes inlet tube is further opened. There occurs again an immediate inrush of hydrogen since the catalyst creates a reduced pressure within the jar due to the conversion of hydrogen and leftover oxygen into water.

If there is no inrush of hydrogen, it means the catalyst is inactive and must be replaced. The jar is left connected to the hydrogen supply for about 5 minutes, then the inlet tube is closed and the jar is placed in the incubator. Catalysis will continue until all the oxygen in the jar has been used up.

The gasPak is now the method of choice for preparing anaerobic jar. The gasPak is commercially available as a disposable envelope containing chemicals, which generate hydrogen and carbon dioxide when water is added. After the inoculated plates are kept in the jar, the gasPak envelope with water added, is placed inside and the lid screwed tight.

Hydrogen and carbon dioxide are liberated and the presence of a cold catalyst in the envelope permits the combination of hydrogen and oxygen to produce an anaerobic environment.

The outstanding feature of the gasPak system is the disposable gas generator envelope, which does away with the need for a vacuum pump and cylinders of compressed gas the operation of the jar is consequently very quick and simple. As the standard gasPak jar is not evacuated before use a relatively large volume of water is formed during catalysis.

An indicator should be used for verifying the anaerobic condition in the jar and for this purpose methylene blue is generally used. When it is placed in an anaerobic environment, it is reduced from its coloured oxidized form to a colourless reduced leuco-compound.

Removal of the culture plates from the jar for microscopic examination is the major disadvantage of any anaerobic jar system. This, of course, results in the exposure of the colonics to oxygen, which is especially hazardous to the anaerobes during their first 48 hours of growth. For this reason, a suitable oxygen-free holding system always should be used in conjunction with anaerobic jars.

The culture plates should be removed from the jar and placed in the oxygen-free holding system. From there they should be removed one by one for rapid microscopic examination of colonies, and then quickly returned to the holding system. Plates never should remain in room air on the open bench.


Making Agar Plates for Bacterial Growth

Agar is medium that cures into a gelatinous form and when mixed with the proper chemicals and nutrients it provides a solid base to grow your bacterial and yeast cultures off of.

These protocols will provide guidance in making the best possible product to provide you with the best possible outcome.

To start we will talk about a bacterial base in which we use LB AGAR.

A common ratio to remember when making your LB AGAR mix is 40g to 1L of water ratio This ratio will make about 80 plates. This protocol is going to walk you through making 10 plates.

10 plates necessitates a 5g LB AGAR powder mixed with 125mL of water.

Put on a pair of Nitrile gloves on for this .

Contamination is critical, as you are providing a platform for bacteria and yeast to grow. They both exist naturally on our skin and in the air, so gloves are a necessity.

Weigh out your LB AGAR on your digital scale.

First turn on your scale, let it zero out, and put a small tray or container to weigh out your LB AGAR on top. Then press the TARE button.

Then slowly add your mix from your tube to the tray until you have reached 6.25 / 6.3g.

Fill your bottle or container that is microwave safe with 125mL water.

Now pour the powder into your water.

Put lid on water and gently shake until you have a consistent yellow fluid. Let the foam settle.

Put cap on container and barely turn it just to hold it in place.

*Note: Do not tighten LID, can possibly make container explode.

Place in microwave for 30 second increments on a normal setting. Stand and watch for boiling as you do not want it to boil over. After all AGAR media has dissolved into a tinted solution, normally 2-3 minutes, you are finished and let cool until it is safe to touch.

While waiting for your media to cool clear off a counter or table and stack plates in columns of 3-5 depending on what you feel comfortable with (Practice grasping the lid of the bottom most unfilled plate and lifting it and all the plates on top of it up).

Ready to pour. So go back to the technique you practiced. Grasp the lid of the bottom most unfilled plate and lift all the plates up. Then pour in the LB Agar until it covers the bottom. Then grasp the next unfilled plate lid in the stack and fill it up.

Try and only add enough to barely fill in the bottom. This will make sure you have more plates!

Let cool and solidify for a few hours or overnight on a table or counter if possible. This lets some of the condensation escape back out before you store them at 4C in your Refrigerator. Store upside down so any condensation doesn&rsquot drip on the plate.

Remember never to freeze plates they will become cracked and distorted.

Plates can be used immediately after they solidify!

Plates should last 2-3 months depending on how much condensation accumulates in the bag and how sterile you were during the preparation.

Below are examples of plates where the LB AGAR was heated properly and not properly


Weigh the quantity of Peptone, Beef Extract, and Sodium Chloride using the weighing scale for 1000 ml of Nutrient Agar Medium as follows:

COMPONENTSQUANTITY (in grams)QUANTITY (in %)
Peptone5 0.5
Beef Extract30.3
Sodium Chloride50.5
Agar-Agar151.5

Put the butter paper on the weighing scale and transfer the required quantity of peptone on to the paper using the spatula. Repeat the step to obtain the required quantity of beef extract, sodium chloride, and Agar-Agar powder.

Note: Use the separate piece of butter paper to avoid the errors in measurements.

Take a clean and dry Conical Flask/ Erlenmeyer flask.

Note: The size of the flask should be at least 1.5 times larger than the quantity of media you are preparing, for e.g. use 1500 or 2000 ml flask to prepare 1000ml of solid medium.

Pour 500 ml of distilled water to the flask and add the weighed quantity of Peptone, Beef Extract, and Sodium Chloride.

Now add the weighed quantity of Agar-Agar to the above solution.

Mix well the content and Heat it with continuous agitation to dissolve the constituents.

Now add more distilled water to the medium and make the volume 1000 ml.

Check the pH of the solution using pH strip, it should be 7.2 ± 0.2. If required, adjust the pH by adding either 1N HCl (acid) or 1N NaOH (base) as per the case.

Mix well the content and apply the Non-absorbent cotton plug to the flask.

Autoclave the content at 121 °C and 15 psi pressure for 15 minutes.

Allow the content to cool down to 40-45 °C and pour in the empty media plates under the strict aseptic atmosphere (preferably in Laminar Air Flow) and allowed it to cool at room temperature.

Use the prepared media plates to inoculate the specimen to be cultured and then place in the incubator at optimum temperature.


Make up nutrient agar as above but using only 900 cm³ of distilled water. Dissolve 20 g of dried skimmed milk in 100 cm³ of distilled water. Sterilize separately. Transfer the milk to the agar aseptically after cooling to 45-50 °C. Dispense aseptically.

Suspend 15 g of nutrient agar in 100 cm³ distilled water. Bring to the boil to dissolve completely. Heat 40 g of soluble starch in 100 cm³ of distilled water to form a suspension. Allow to cool and then mix with the nutrient agar solution. Dispense and sterilize.


Exercise: Transformation of Bacteria with RE Identified Plasmids

  1. Each group retrieves the 2 miniprepped plasmids from the previous week in the freezer and allow to thaw on ice.
  2. Bring 2 agar plates to room temperature
    • 2 plates will contain antibiotic, X-Gal and arabinose
    1. For each plasmid, obtain 250μl of transformation buffer (50mM CaCl2) in microfuge tubes and place on ice for 10 minutes
    2. Take an inoculating loop and remove a single colony of bacteria from a freshly streaked plate grown overnight
    3. Swirl bacteria in each tube containing transforming solution to distribute bacteria throughout solution
    4. Pipette 5 μl of plasmid into the tube and incubate on ice for 10 minutes
    5. During this incubation, flip the warmed plates and label them with your group names.
    6. Place transformation tubes into 42°C heatblock for 1 minute to heat shock the cells
    7. Add 500μl fresh SOC media (or LB) and incubate at 37°C for 15 minutes.
    8. Pipette 150μl of transformation solution onto each plate and spread across the plate.
    9. Turn plates agar side up and place them into 37°C incubator overnight. (your instructor will retrieve them and place them into refrigerator)

    Method for preparation of LB agar plates with Ag+? - Biology

    1 - Coliphage lambda DNA is a widely used vector for recombinant DNA. The middle third of its 48,000 bp contains no genes required for lytic growth and is, therefore, replaceable. The usual recombinant lambda DNA contains 80% lambda vector DNA and 20% insert, as compared to a usual cosmid DNA that contains 10% vector and 90% insert.

    Wild-type lambda is not very lytic, compared to coliphages T4 or T7. Recombinant lambda phage are usually less lytic than wild-type. We recommend growing recombinant lambda phage on large (150-mm diameter) plates, rather than in liquid culture. Beginners, in particular, may not be aware sufficiently that in liquid culture E. coli can easily overgrow phage lambda, rather than the desired vice versa .

    2 - Traditionally, the E. coli host for lambda is grown on NZY medium. This medium is not as rich as LB (and many other) medium, and E. coli growth is slower, making it even less likely that E. coli will overgrow the phage.

    The lambda receptor on the bacterial cell outer surface is part of the maltose-usage pathway. To induce the receptor to high levels (10 2 -10 3 receptors per cell), maltose is added to a final concentration of 0.2% in the medium.

    When growing lambda on plates to prepare DNA (as opposed to picking plaques or titering), agarose is substituted for agar. Agarose is much more expensive than agar, but does not contain impurities that inhibit enzymes. Lambda DNA prepared from agar plates may not be digestible by restriction enzymes because of the presence of inhibitors that have leeched from the agar.

    3 - The host for lambda is E. coli C600 and its subsequent variants. To avoid complications, the E. coli C600 strain is usually restriction-negative: r-m- or r-m+ .

    For our purposes, we find it useful to start with a fresh E. coli colony (on NZY or LB plates). The colony is picked into 5 ml of NZY medium plus 0.2% maltose and placed on a wheel at 37°C for approximately 4 hr. (This time will be much longer if the colony is from an old plate or has been stored at 4°C or frozen.) Using sterile conditions, the culture is transferred to a sterile 15-ml conical tube. The E. coli are pelleted by centrifugation in an SS34 rotor with adaptors in an RC-5B centrifuge (or equivalent) for 8 min at 4 K rpm (or equivalent). The medium is poured off gently. The bacterial pellet is resuspended in 4-6 ml of 0.01 M MgSO4 by pipetting up and down (using sterile conditions), NOT by vortexing. The concentration of bacteria is adjusted to yield an "optical density" (really, light scattering) of 2.0 at wavelength 600 nm. E. coli prepared in this way can be stored at room tempera ture and used for up to one week. To resuspend the E. coli , gently pipet the cells. DO NOT VORTEX. Cells prepared in this manner are employed at values of 10 µl per small or 20 µl per large plate.

    4 - At times, we shall refer to the procedures published in Sambrook, Fritsch, and Maniatis, in "Molecular Cloning: a laboratory manual", pp. 2.60-2.79, second edi tion, 1991, as the "CSH" procedure or the "CSH" book.

    One common source of confusion when comparing methods for growing lambda concerns "SM buffer" and whether or not it contains gelatin. In our protocol, "SM buffer" refers solely to Tris buffer with magnesium salt and NaCl. If gelatin is to be added, we state so explicitly in the few cases where we use it.

    5 - The start of any procedure for preparing lambda DNA is to pick a single, well -isolated plaque. (A lambda plaque contains phage and lysed bacteria and appears relatively "clear" against the bacterial lawn.) For the Olson frozen lambda stocks, we use sterile technique to scrape a small amount (approximately a toothpick-full) of frozen stock into 1 ml of SM buffer plus gelatin. (The gelatin helps stabilize the phage.) From this one ml, we make 10-fold dilutions (in SM buffer plus gelatin) through 10-4. In general, we plate 1, 10, and 100 µl of the 10 -3 and 10-4 dilutions.

    There is significant preparation time. NZY agar plates should be placed at 37°C, so that they are warm when used. NZY top agar (3 ml per plate) should be melted and placed at 50°C (just warm enough to keep it liquid, and not so hot as to kill the E. coli ). E. coli (10 µl) and 0.2 ml SM buffer (no gelatin) should be mixed in a small tube, one tube per agar plate.

    Phage are added to the E. coli and SM buffer, and the tubes are placed on the 37°C wheel for 20-30 min. The kinetics of lambda attachment to its receptor are fairly slow the 20 min incubation allows time for this reaction to occur. Add the 3 ml of top agar to the mixture of phage and bacteria. Vortex for 10 sec. Pour onto an agar plate. Rock the plate by hand to reach an even distribution of top agar. Allow the top agar to solidify at room temperature (approximately 10 min). Place the inverted plates at 37°C overnight. The next day, at least one plate should have well-isolated plaques. If there are too many plaques, start over with more dilution. If there are too few plaques, start over with less dilution.

    For picking plaques, we find the following procedure simple and straightforward. Autoclave inverted Pasteur pipets (without cotton plugs). Holding the narrow end, punch out a plaque with the (relatively) wide-mouth end. The agar stays in the pipet and can be expelled into a sterile eppendorf tube by a wrist flick. Add 0.5 ml of SM buffer (no gelatin) and 2 drops of chloroform (the chloroform stops the E. coli from growing). To elute the phage, allow the eppendorf tube to stand for 2-4 hr at room temperature or at 4°C overnight.

    6 - A phage stock is made from the isolated plaque. Again, NZY agar plates are at 37°C NZY top agar is at 50°C 0.2 ml of SM buffer plus 10µl of E. coli are in small tubes. Add 5-20 µl (or more) of phage, depending on plaque size, to the E. coli and SM buffer. Place on a 37°C wheel for 20-30 min. Add NZY top agar and vor tex for 10 sec. Pour onto a warm NZY agar plate. After the top agar hardens (approximately 10 min), invert the plate and place at 37°C overnight.

    The next morning the plates should show a fully infected, disrupted E. coli lawn. If you see individual plaques, the inoculum was too low. Toss the plates, and repeat the infection with a larger volume of the plaque. If the infection looks confluent, add 5 ml of sterile SM buffer (no gelatin). Make sure that the plate is completely covered with buffer. Gently rotate or rock the plates at room temperature for 1-2 hr. Harvest the buffer (using a sterile pipet) into a sterile conical 15-ml tube. The "milky" appearance is caused by the presence of bacterial debris. Add 0.2 ml of chloroform and vortex for 10 sec. Spin the tube in an SS34 rotor with adaptors, RC-5B centrifuge (or equivalent), at 4 K rpm for 8 min. Decant the clear yellow supernatant into a fresh, sterile, 15-ml conical tube. Add a drop of chloroform (the chloroform is to stop the growth of residual E. coli ) and store at 4°C. These stocks are stable for several months at 4°C. Frozen (-70°C) stocks can be made by mixing 0.1 ml DMSO with 1.4 ml of lambda phage stock.

    7 - The phage stocks of Olson lambda are usually in the range 10 5-106 pfu (plaque-forming units) per µl. These numbers are lower than those given in CSH, pg. 2.65 the Olson lambda phage grow very poorly. Our stocks have been as low as 2x10 3 and as high as 8x107 pfu per µl. The general correlation is that phage that pro duce small plaques tend to yield stocks below 105 pfu per µl phage that produce (relatively) large plaques tend to yield stocks over 10 6 pfu per µl but the correlation is not absolute.

    The lambda stocks are diluted ten-fold (as appropriate: 10 -2, 10-3, etc .) in SM buffer plus gelatin. NZY agar plates are at 37°C. NZY top agar is melted, aliquoted, and maintained at 50°C. 0.2 ml of SM buffer plus 10 µl of E. coli are in small tubes. Appropriate amounts (e.g., 1, 10, 100 µl) of each appropriate dilution are added to the bacteria. The tubes are placed on the 37°C wheel for 20-30 min. 3 ml of top agar are added to the phage and bacteria the tube is vortexed for 10 sec, and poured onto a warm agar plate. Distribute the top agar evenly on the plate while rocking (rotating, swirling) by hand. Allow the top agar to harden at room temperature (approximately 10 min). Invert the plates and place at 37°C overnight.

    The next morning look at the the plaques. They are usually of a reasonable size to count. However, if the plaques are very small (which does occur with the Olson lambdas), leave the plates at 37°C and check the size of the plaques hourly. Count the plaques on plates where the number is between 50 and 300. A mini mum of 100 is needed for reasonable statistical reliability a plate with over 200 plaques may yield an incorrectly low number, as plaques overlap. How much accurracy you need is up to you. Calculate the titer of the stocks.

    8 - With the preparation of titered phage stocks derived from a single plaque, all the reagents are in hard to prepare lambda DNA. In addition to titered phage stocks, you need large (150-mm) NZY agarose plates, NZY top agarose, and E. coli C600 in 0.01 M magnesium sulfate. We switch from agar to (the much more ex pensive) agarose to avoid the umwanted impurities present in agar. We usually use two 150-mm plates for growth of each original phage plaque. The large NZY agarose plates are equilibrated at 37°C. NZY top agarose is at 50°C. Place 0.2 ml of SM buffer in a small tube and add 20 µl of E. coli (double the amount for a small plate). For poor-growing lambda, such as the Olson recombinants, we add 50,000 pfu to the tube. However, this number can vary greatly: higher for very poor -growing phage and much lower for phage which grow well. You may have to test several phage inputs. The goal is to achieve confluent lysis, maximum phage pro duction, without blowing away the E. coli prematurely. Add the lambda to the bacteria and SM buffer and place the tube on the 37°C wheel for 20-30 min. Add 5 ml (an increased amount for the large plates) of top agarose vortex for 10 sec, and pour onto the warm 150-mm plate. Rotate (rock, swirl) the plate by hand to achieve an even distribution of top agarose. Allow the top agarose to harden (approximately 10 min) and invert the plate at 37°C overnight.

    The next morning the plates should show confluent lysis. If individual plaques are visible, you may not achieve a usable amount of phage DNA. Start over at a higher input. We find that two confluently-lysed large plates will yield adequate amounts of DNA, even for poorly-growing lambdas.

    9 - Add 8 ml of SM buffer to each plate. Make sure the buffer covers the entire plate. Sterile conditions are no longer necessary, but neatness always counts. Rotate gently (or rock gently) at room temperature for 1 to 2 hr. The phage will elute into the buffer.

    Decant (pipette) the buffer into a 15-ml conical tube (or equivalent): 5-6 ml/plate. Two large plates of the same phage are combined, as appropriate. Centrifuge the tube in an SS34 rotor with adaptors, RC-5B centrifuge (or equivalent) for 8 min at 4 K rpm (or equivalent). The debris will pellet, and most of the phage will reamain in the supernatant. Decant the supernatant (9-12 ml) to an Oak Ridge tube.

    10 - In this next , crucial step, DNase I is added to digest contaminating E. coli DNA. Lambda DNA, inside the phage particle, is protected from digestion. The integrity of the phage requires the presence of magnesium ions, and DNase activity requires magnesium ions, which are provided in SM buffer. Add 0.1 ml of DNase I (100 µg per ml RNase-free) and mix gently. Place the tube at 37°C for 1-2 hr. (No attempt is made to remove RNA, and, in fact, RNA is used as a carrier for the DNA.)

    11 - Inactivate DNase I activity and disrupt the phage particles by the additions of 0.5 ml of 0.5 M EDTA, pH 8, and 0.5 ml of 10% SDS. Mix gently. (The removal of magnesium ions by the EDTA also causes the polysomes and ribosomes to fall apart. This helps during phage DNA purification.)

    12 - To remove protein and SDS, add 10 ml of phenol (previously equilibrated with TE buffer). Vortex hard for 20 sec. Centrifuge in an SS34 rotor, RC-5B centrifuge (or equivalent), for 10 min at 10 K rpm at 10°C. Decant the upper, aqueous phase to a fresh Oak Ridge tube. (We use an inverted 10 ml pipette without cot ton plug to transfer.) Add 10 ml of chloroform. Vortex hard for 20 sec. Centri fuge in an SS34, RC-5B centrifuge (or equivalent), for 10 min at 10 K rpm at 10°C. Decant the upper, aqueous phase to a fresh Oak Ridge tube (again we use an in verted 10 ml pipette). Note the volume transferred (usually around 10 ml).

    13 - Concentrate the DNA, and remove traces of phenol and chloroform, by alcohol precipitation. Add two-volumes of cold (95-100%) ethanol. Vortex for 10 sec to mix. Centrifuge in an SS34 rotor, RC-5B centrifuge (or equivalent), for at least 30 min at 12 K rpm at 10°C. Gently decant the supernatant. (We pour the alcohol off carefully while watching that the precipitate does not move.) To remove ex cess salt, gently add 10 ml of cold 70% ethanol. Do not disturb the nucleic acid precipitate. Centrifuge for 10 min (or more) at 12 K rpm at 10°C. Gently decant the supernatant. Add 10 ml of cold (95-100%) ethanol. Centrifuge for 10 min utes at 12 K rpm at 10°C. Gently decant the supernatant. Drain briefly and air dry briefly. Our experience with the Olson lambda DNAs is that the amount of precipitate is highly variable. Most of the precipitate is RNA.

    14 - Dissolve the precipitate in 2.5 ml of TE. This process may take longer than 1 hr at room temperature. Add 0.1 ml RNase (1 mg/ml, DNase free) and mix gently. (At this step, the contaminating RNA is removed by digestion). Place at 37°C for 1-2 hr. Add 0.1 ml of proteinase K (1 mg/ml, nuclease-free) and incubate at 37°C for 1-2 hr. Contaminating protein, including RNase and proteinase K itself, is re moved by digestion. Cool to room temperature.

    15 - Add 2.5 ml of phenol:chloroform (1:1 the phenol has previously been equili brated with TE buffer). Vortex hard for 10 sec. Centrifuge for 10 min at 10 K rpm, 10°C, in an SS34, RC-5B centrifuge (or equivalent). Decant the upper, aque ous layer to a Centricon-30 (Amicon Co.).

    16 - The Centricon takes the place of alcohol precipitation and/or dialysis to change buffer and concentrate the DNA. The Centricon works annoyingly slowly, but the recoveries are excellent. Read the manufacturer's (Amicon Co.) instruc tions. The plastic that composes the Centricon is resistant to traces of phenol and chloroform but is easily damaged by isoamyl alcohol. Many recipies and commer cially available phenol:chloroform add isoamyl alcohol as an anti-foam reagent. Do not use those recipes/products, or do not use the Centricon.

    We use a Centricon-30 spun at 5 K rpm in an SS34, RC-5B centrifuge at 10°C. Other rotor/centrifuge combinations may have a different maximum speed check Amicon's specifications. The choice of 10°C, rather than 4°C, is to avoid icing-up.

    Under these conditions, a 2.5 ml sample will pass through the filter in about 30-60 min. We load the sample completely, do three 2.5 ml TE washes, and then spin for an additional 2 hr to reach minimum volume (usually 50-60 µl).

    17 - For the Olson lambda DNAs, we use 7 µl of the DNA to double-digest and run on a 1% agarose gel in 0.5xTBE. If all has gone well, the restriction enzyme cleav age pattern matches Olson's.

    Standard reagents for DNA isolation and manipulation include:

    TE buffer : 0.01 M Tris, 0.001 M EDTA, pH 8.0 usually diluted from a 100x stock: 1.0 M Tris, 0.1 M EDTA, pH 8.0.

    0.5 M EDTA, pH 8.0: EDTA is sold as the disodium salt. In solution, Na 2EDTA has a pH near 5 and a saturation limit around 0.2 M . To achieve 0.5 M , NaOH needs to be added in small aliquots.

    10% SDS : 10 g of sodium dodecyl sulfate dissolved in water to a final volume of 100 ml.

    Ethyl alcohol : 95 or 100% and 70%, both at -20°C.

    Phenol : Standard commercial phenol is quite dirty by molecular biology standards and must not be used. Several companies sell re-distilled phenol of "nucleic acid" or "molecular biology" grade. We are using Amresco's saturated phenol, which is a liquid, and comes with a small bottle of TE buffer to be added to the phenol. We want buffer-saturated phenol. Add the buffer to the phenol and shake. Allow to stand overnight in a refrigerator. Use an amber bottle to avoid light. We want to avoid oxidation of the phenol to produce quinones, which are highly reactive and detrimental to DNA. A reminder: a saturated solution cannot be created in an hour. At saturation, the upper phase is excess buffer.

    Chloroform (CHCl3) : any brand at A.C.S. purity is fine.

    Phenol: Chloroform (1:1) : Mix one volume of buffer-saturated phenol with the same volume of CHCl3 in an amber bottle. Shake and place in the refrigerator overnight. The upper phase is excess buffer. We make our own mixture, because Amresco's phenol: chloroform contains isoamyl alcohol as an anti-foam reagent, and isoamyl alcohol dissolves the plastic Centricon.

    For this procedure, the following reagents are required:

    0.01 M MgSO4 : used to resuspend E. coli C600. We use A.C.S. grade MgSO4ܭH 2O. Sterilize by autoclaving or filtration.

    DNase I (free of RNase) : Several companies sell this reagent. Our stock solu tion is 100 µg/ml in 0.01 M Tris, pH 8.0. We purchase DNAse I from Boehringer -Mannheim. The stock is aliquoted, frozen (-20°C), and thawed as needed. (no EDTA) Careful: DNAse I is very heat labile and is easily inactivated at room tem perature.

    RNAse (free of DNAse): We purchase this enzyme from Boehringer-Mannheim. (Even the smallest amount of contaminating DNAse will degrade your precious DNA.) The RNAse is aliquoted and frozen (-20°C).

    Specialized reagents for growing coliphage lambda:

    • NZ-amine (also known as "casein enzymatic hydrolysate" e.g., Sigma cat.# C -0626) 10 g
    • Yeast extract 5 g
    • NaCl 5 g
    • MgSO4ܭH2O 2 g

    Sterilize by autoclaving. For agar plates (100-mm plates used for counting or isolating plaques), add 15 g of agar per liter before autoclaving. For top agar, add 7 g of agar per liter before autoclaving. For ease of handling, we prepare top agar in 100-200 ml batches, rather than 1 l batches. For large (150-mm) agarose plates, used when growing lambda to prepare DNA, add 15 g of agarose per liter before autoclaving top agarose requires 7 g of agarose per liter before autoclav ing. Unless you have an unlimited supply budget, you should use agarose ony where essential. We make both NZY-agar and NZY-agarose plates and top media to save money.

    Maltose (20%) : Dissolve 20 g of maltose in a final volume of 100 ml. Sterilize by filtration. Maltose, as all sugars, tends to become caramel upon autoclaving.

    (The CSH book uses the nomenclature "Tris·HCl", which is not technically correct. We start with Tris·OH and add HCl to pH 7.5. Therefore, our Tris buffers do not have additional sodium ions. Incidentally, Tris·OH is half of the cost of Tris·HCl.)


    • Luria-Bertani broth (though this name is very widely used)
    • Luria broth
    • Lennox broth

    The recipe for LB was formulated by Giuseppe Bertani and published in 1951. Over the years the acronym has been widely misconstrued. In the Postscript to his 2004 paper, “Lysogeny at Mid-Twentieth Century: P1, P2, and Other Experimental Systems”, Giuseppe Bertani clarified the original meaning of the acronym:

    “My first paper on lysogeny, describing the modified single-burst experiment and the isolation of P1, P2, and P3, also contained the formula of the LB medium which I had concocted in order to optimize Shigella growth and plaque formation. Its use has since become very popular. The acronym has been variously interpreted, perhaps flatteringly, but incorrectly, as Luria broth, Lennox broth, or Luria Bertani medium. For the historical record, the abbreviation LB was intended to stand for “lysogeny broth.” (5, page 598).


    Exercise: Transformation of Bacteria with RE Identified Plasmids

    1. Retrieve miniprepped plasmid from freezer and allow to thaw on ice.
    2. Bring 2 agar plates to room temperature
      • 1 plate will contain antibiotic, X-Gal and arabinose
      • 1 plate will contain only antibiotic
      1. Pipette 250μl of transformation buffer (20mM CaCl2) into a microfuge tube onto ice for 10 minutes
      2. Take an inoculating loop and remove a single colony of bacteria from a freshly streaked plate grown overnight
      3. Swirl bacteria in tube with transforming solution to distribute bacteria throughout solution
      4. Pipette 5 μl of plasmid into the tube and incubate on ice for 10 minutes
      5. During this incubation, flip the warmed plates and label them with your group names. Note which one has X-Gal and Arabinose.
      6. Place transformation tubes into 42°C heatblock for 1 minute to heat shock the cells
      7. Add 250μl fresh LB and incubate at 37°C for 15 minutes.
      8. Pipette 200μl of transformation solution onto each plate and spread across the plate.
      9. Turn plates agar side up and place them into 37°C incubator overnight. (your instructor will retrieve them and place them into refrigerator)

      Synthesis, Characterization, and Evaluation of the Antibacterial Activity of Allophylus serratus Leaf and Leaf Derived Callus Extracts Mediated Silver Nanoparticles

      Allophylus serratus mediated silver nanoparticles biosynthesis, characterization, and antimicrobial activity were described. The synthesis of silver nanoparticles was confirmed by visual observation: UV-Vis spectrum, X-ray diffraction (XRD), Scanning Electron Microscopy (SEM), Energy Dispersive Spectroscopy (EDS), and Fourier Transform Infra-Red (FTIR). UV-Vis spectroscopy studies showed that the absorption spectra of synthesized silver nanoparticles from leaf and callus extracts had absorbance peak range of 440 nm and 445 nm, respectively. The X-RD pattern revealed the presence of crystalline, dominantly spherical silver nanoparticles in the sample having size ranging from 42 to 50 nm. The XRD peaks 38.2°, 44.1°, 64.1°, and 77.0° for leaf extract and 38.1°, 44.3°, 64.5°, 77.5°, and 81.33° for callus extract can be assigned the plane of silver crystals (111), (200), (220), and (311), respectively, and indicate that the silver nanoparticles are face-centered, cubic, and crystalline in nature. SEM and EDS analysis also confirmed the presence of silver nanoparticles. The FTIR results showed the presence of some biomolecules in extracts that act as reducing and capping agent for silver nanoparticles biosynthesis. The synthesized silver nanoparticles showed significant antibacterial activity against Klebsiella pneumoniae and Pseudomonas aeruginosa.

      1. Introduction

      Nanotechnology is a field of science which deals with production, manipulation, and use of nanomaterials ranging in size from 1 to 100 nanometers. Nanomaterials have novel and enhanced useful characteristics due to their size, distribution, and morphology in comparison to the larger particles of the mass material that they have been prepared from (Wildenberg 2005). Due to their enhanced or new properties, novel applications of nanomaterials and nanoparticles are growing rapidly on various fields such as electronic, magnetic, optoelectronics, and information storage (Sun et al. 2002 Yin et al. 2003). They are also broadly applied in shampoos, detergents, soaps, cosmetics, toothpastes, and medical and pharmaceutical products (Bhattacharya and Murkherjee 2008 Bhumkar et al. 2007). Nanoparticles also have been proven to have antimicrobial activity and used in the field of medicine. They are also used in cosmetics, healthcare, biomedical, drug-gene delivery, food and feed, environment, mechanics, chemical industries, optics, electronics, space industries, energy science, catalysis, single electron transistors, light emitters, nonlinear optical devices, and photo-electrochemical applications [1].

      Among nanomaterials and nanoparticles used for all the above listed purposes, the metallic nanoparticles are considered as the most promising due to their remarkable antimicrobial properties. This is of great interest for researchers due to the development of resistant strains of microbes against antibiotics [2]. Compared to other heavy metal nanoparticles, silver nanoparticles are very important because of their unique properties such as good conductivity, chemical stability, catalytic and most important antibacterial, and antiviral, antifungal, and anti-inflammatory activities. They can be incorporated into composite fibers, food industry, cryogenic superconducting materials, electronic components, and cosmetic products [3, 4]. Silver nanoparticles are also applied in textile, home water purification systems, medical devices, cosmetics, electronics, and household appliances [5]. They are also being used in cancer diagnosis and treatment [6, 7].

      Recently, synthesis of silver nanoparticles is attracting the attention of scientific community due to their wide range of applications. There are different chemical, physical, and biological methods of synthesizing nanoparticles. Most of the chemical and physical methods of silver nanoparticles synthesis are expensive and involve use of toxic and hazardous chemicals which are potentially harmful to the environment and responsible for various biological risks. Biological or green synthesis of nanoparticles by using biological materials such as plant extract, plant biomass, or microorganisms is environmentally friendly, is safe, and helps to reduce the consequence of chemical methods of nanoparticles synthesis [8, 9].

      Some of the biological methods of silver nanoparticles synthesis involve very complex procedures. For example microbial mediated synthesis of silver nanoparticles is not industrially practical due to requirement of high aseptic and maintenance conditions [1]. Synthesis of silver nanoparticles using various plants materials and their extracts is simple way and beneficial over other biological synthesis processes which involve very complex procedures (Sastry et al. 2003) [10].

      The use of plants for the synthesis of silver nanoparticles has drawn attention not only due to its nonpathogenic, environmental friendly, and economical protocol but also because of being a simple and single step rapid technique. A large number of plants and their respective portions are reported to facilitate silver nanoparticles synthesis. Plant extracts from various plants such as Alternanthera dentata [11] Acorus calamus [12] Boerhaavia diffusa [13] Tea [14] Sesuvium portulacastrum [15] Tribulus terrestris [16] Cocos nucifera [17] A. indicum (Ashok et al. 2015) Ziziphora tenuior [18] Ficus carica [19] Cymbopogon citratus [20] Acalypha indica [21] Chenopodium album [22] Cocos nucifera [17] Pistacia atlantica [23] Cymbopogon citratus [24], and so forth have been reported for the synthesis of silver nanoparticles.

      In our present study, we investigated synthesis of silver nanoparticles from leaf and leaf derived callus extracts of Allophylus serratus, their characterization, and evaluation of antibacterial activity. This work is the first report on synthesis of silver nanoparticles using this plant which is an additional confirmation of previous reports on biological synthesis of silver nanoparticles using plant leaf and callus extracts and their antimicrobial activity.

      2. Materials and Methods

      2.1. Materials

      Healthy fresh leaves of Allophylus serratus were collected from Andhra University campus in month of January 2016. The leaves were identified and authenticated by Dr. Bodaih Padal, taxonomist, Department of Botany, Andhra University, Visakhapatnam. All chemicals and reagents used in this study silver nitrate AgNO3 (99.98%), 6-Benzylaminopurine (BAP), 1-Naphthaleneacetic acid (NAA) ethanol (C2H6O), sodium hypochlorite (NaClO), mercuric chloride (HgCl2), Murashige and Skoog media (MS media), Luria Bertani broth (LB broth), and Luria Bertani agar (LB agar) were analytical grade (AR) and purchased from HiMedia, India. Double distilled water was used throughout the experiment. The bacterial strains, namely, Bacillus subtilis, Staphylococcus aureus, Klebsiella pneumoniae, and Pseudomonas aeruginosa, were obtained from Department of Microbiology, Andhra University, Visakhapatnam, India. All solutions were freshly prepared using double distilled water and kept in the dark to avoid any photochemical reactions. All glassware used in experimental procedures was washed thoroughly with double distilled water and dried using hot air oven before use.

      2.2. Callus Induction and Proliferation

      Callus induction of Allophylus serratus was achieved from leaf explants on MS media [25]. The PH of the media was adjusted to 5.8 and then autoclaved at 121°C for 15 min under 103.42 KPa pressure. In laminar hood, plant growth regulators BAP (3 g/L) NAA (0.5 g/L) were added by filter sterilization. The leaf explants were surface sterilized by using 70% ethanol for 1 minute, 2% sodium hypochlorite for 3-4 minutes, and 0.1% mercuric chloride solution for 3 minutes. Finally the leaf explants were washed thoroughly with sterile double distilled water and inoculated in the MS medium. The cultures were kept in dark at 25°C. The induction of the callus was found within 2 weeks. The callus was subcultured every 21 days several times and finally a mass of calli was harvested after 40 days.

      2.3. Preparation of Leaf and Leaf Derived Callus Extract

      Leaves of Allophylus serratus were washed, shade dried for one week, and ground to powder. Callus induced from leaves on MS medium supplemented with 3 mg/L BAP and 0.5 mg/L NAA was grown for 40 days and collected and dried in oven at 40°C. Then the calli were ground to powder using mortar and pestle. These powders of leaves and callus were used for the synthesis of silver nanoparticles (Figure 1).


      Watch the video: LB Plate Preparation (August 2022).