1: Media Preparation - Biology

Learning Objectives

  • Understand how to make media, how to sterilize it, and how to distribute it in different formats.
  • Produce TSA plates, TSA slants, and TSB which will be used in subsequent lab periods.
  • Understand the basics of an autoclave and how it sterilizes, including parameters.

Bacteria and fungi are grown on or in microbiological media of various types. The medium that is used to culture the microorganism depends on the microorganism that one is trying to isolate or identify. Different nutrients may be added to the medium, making it higher in protein or in sugar. Various pH indicators are often added for differentiation of microbes based on their biochemical reactions: the indicators may turn one color when slightly acidic, another color when slightly basic. Other added ingredients may be growth factors, (ce{NaCl}), and pH buffers which keep the medium from straying too far from neutral as the microbes metabolize.

In this exercise, you will make all-purpose media called trypticase soy broth and trypticase soy agar. These 2 media----one a liquid and the other a solid---are the exact same formula save for the addition of agar agar (really- agar agar), an extract from the cell walls of red algae.

The old way to make media was by the cookbook method--- adding every ingredient bit by bit. The only time that is done today is when making a special medium to grow a certain finicky organism, where particular growth factors, nutrients, vitamins, and so on, have to be added in certain amounts. This medium is called a chemically defined medium (synthetic). Fortunately, the most common bacteria that we want to grow will do nicely with media that we commonly use in lab. Some of our media is bought, but most is produced in the prep area behind the lab. Since this type of medium has some unknown ingredients, or sometimes unknown quantities it is called complex media.

It is really very simple to make complex media these days:

  • rehydrate the powder form of the medium
  • stir and boil the agar medium to get the agar powder dissolved (if making an agar medium rather than a broth medium)
  • distribute the medium into tubes
  • autoclave to sterilize the tube media
  • autoclave the agar medium for plate production and then pour into sterile petri dishes


When microbiological media has been made, it still has to be sterilized because of microbial contamination from air, glassware, hands, etc. Within a few hours there will be thousands of bacteria reproducing in the media so it has to be sterilized quickly before the microbes start using the nutrients up. The sterilization process is a 100% kill, and guarantees that the medium will stay sterile UNLESS exposed to contaminants by less than adequate aseptic technique to exposure to air.

Media sterilization is carried out with the autoclave, basically a huge steam cooker. Steam enters into a jacket surrounding the chamber. When the pressure from the steam is at a certain point in the jacket, a valve allows the steam to enter the chamber. The pressure will go up over 15 pounds per square inch (psi): at this point the timer begins to count down--- usually for 15 minutes, depending on the type of media. The high pressure in a closed container allows the temperature to go above the highest temperature one could get by just boiling, around 121⁰C. Therefore, the parameters for sterilization with an autoclave are 121⁰C at >15 psi for 15 minutes. Fifteen minutes is the thermal death time for most organisms (except some really hardy sporeformers).

The prepared media is distributed in different ways, depending on the form one is making. Broths and agar deeps are dispensed into tubes and then sterilized. Agar slant tubes are sterilized and then the rack is tilted to allow the agar to solidify in a slanted fashion. Agar medium to be be poured into plates is sterilized in a flask, and then poured afterward. Not all media or solutions can be sterilized via an autoclave. Certain high-protein solutions such as urea, vaccines, and serum will denature in the extreme heat, and so they may have to be filter-sterilized without heat. You will be making slant and broth media, but not plate media in this lab.


  • 2 plastic weigh boats
  • 1 test tube rack
  • 1-1 liter Erlenmeyer flask
  • 1 pipet pump
  • 1 graduated cylinder several nonsterile glass10 ml pipets
  • 1 spatula
  • 28 medium, nonsterile test tubes
  • 1 jar agar powder 15 green caps
  • 1 jar nutrient trypticase soy broth powder
  • 15 yellow caps
  • 1 magnetic stir bar
  • 1 pipet disposal jar


Refer to the diagram below for the entire production:

  1. Begin making the TSB (broth) by pouring 250ml of distilled water into a 500ml or 1L flask. Put in the stir bar and turn on the stir plate so that the surface is just disturbed. Add 3.25 grams of the TSB powder to this flask and allow it to dissolve (will happen quickly). No heat need be applied at this stage.
  2. Once the powder is dissolved, pipet out 5ml green cap.

Green caps are always used for TSB.

  1. With the remaining solution (about 100ml) still stirring, add 2 grams of agar powder.
  2. The next step will require you to apply heat to the mixture. Before you do this, however, you should be aware that agar has a strong tendency to boil over when it reaches 100⁰C. Someone in your group should be watching the flask at all times once you see steam coming off of it. At the first sign that the mix is near boiling, REMOVE it from the hot plate (paper towels around the flask neck). DO NOT simply turn off the heat, letting the flask sit there. The metal plate retains a significant amount of heat, and turning off the heat will not prevent the flask from boiling over. Folded paper towels allow you to grasp the flask neck tightly, yet not burn your hand.
  3. Have you read step 4? OK, then you can turn on the heat to setting 9 (not High). Make sure that the magnetic bar is stirring the solution.
  4. Upon boiling, the agar dissolves, it will turn clear, deeper tan. Remove it from the heat and pipet out 5ml aliquots into 15 tubes for slants (will not be BE slants until removed from autoclave and tilted to the side to solidify). Cover the slant tubes with yellow caps. THE REST OF THE AGAR MEDIUM IN THE FLASK WILL BE POURED INTO 1 LARGE FLASK FOR THE CLASS.

From this point on, yellow caps will be used for nutrient agar slants.


If the agar solidifies in the tip of the pipet, dispose of the pipet in the pipet jar and get another one. To prevent this from happening, either pipet out all the tubes at the same time, or leave the pipet in the flask of melted agar.

  1. Place all of the tubes you have pipetted out in the plastic autoclave racks on the instructor's table as well as the remaining of your melted agar. All agar slants go in one rack, broths in another rack, etc.
  2. Dispose of your used pipets in the pipet holder. These glass pipets are reusable, so don't throw them in the trash.


  1. What is a complex medium?
  2. Why are pH buffers added to the growth media for microbes?
  3. How can the temperature in the autoclave go above boiling temperature of 212 F?
  4. Why do you have boil the agar solution BEFORE dispensing it into tubes?
  5. At what temperature does agar solidify?

Culture of Microorganisms: 5 Steps

The following points highlight the five main steps for culture of microorganisms. The steps are: 1. Preparation of Media 2. Adjustment of pH of Media 3. Preparation of Stabs and Slants 4. Pouring of Plates 5. Inoculation of Bacteria in Nutrient Slants and Agar Plates.

Step # 1. Preparation of Media:

In preparing a culture medium for any microorganism, the primary goal is to provide a balanced mixture of the required nutrients, at concentrations that will permit good growth. No ingredient should be given in excess because many nutrients become growth inhibitory or toxic as the concentration is raised.

A medium composed entirely of chemically defined nutrients is termed as synthetic medium. One that contains ingredients of unknown chemical composition is termed a complex medium. For different purposes in laboratory the media is used either ‘solid’ or ‘liquid’.

A. Liquid or Broth Media:

Nutrient broth is the basis of most media used in the study of different types of microbes. It is one of the most important liquid media used for bacteriological purposes.

(i) Flask – 1000 ml capacity.

(ii) Graduated cylinder – 500 ml.

(iii) Beef extract, bactopeptone, distilled water.

(iv) Balance and weight box.

(v) pH paper, comparator block, 0.1 (N) NaOH and 0.1 (N) HCl soln.

To make 300ml of nutrient broth: –

0.9 gm. of beef extract and 1.5 gms of bactopeptone are weighed separately and taken in a coni­cal flask. Then 300 ml dist. water is added and the ingredients are mixed thoroughly. The pH is adjust­ed by adding a little alkali (NaOH soln.). The flask is then plugged with cotton wool and autoclaved at 15 lbs. pressure for 15 minutes.

2. Potato-dextrose Broth:

It is a type of semisynthetic medi­um. This is very often used for the growth medium of fungi which grow better in potato-dextrose broth than in the nutrient broth.

Freshly peeled potato – 400 gms. (40%)

(ii) Graduated cylinder – 250 ml.

(iii) Potato, Dextrose and distilled water.

(iv) Other requirements are as in the previous broth preparation.

To make 200 ml of broth:

80 gms of freshly peeled potato and 5 gms of Dextrose are weighed and taken in a conical flask. 200 ml of dist. water is poured into the flask and the ingredients are mixed thoroughly by a glass rod. (Actually the peeled potato is boiled in flask using 100 ml water for 10 min and the extract is taken by decanting). The flask is plugged and autoclaved at 15 lbs. pressure for 15 min.

B. Solid or Agar Medium:

Nutrient agar is an important medium for bacteriological purpose. It is simply nutrient broth solidified by the addition of agar. Due to its solid consistency, this medium serves as a good device for the culture of bacteria on a solid surface.

Bacteriological peptone – 5 gms. (0.5%)

ii) Graduated cylinder – 500 ml.

iii) Beef extract, bactopeptone, agar, and distilled water.

iv) Balance and weight box.

v) pH paper, comparator block 0.1(N) HCl and 0.1 (N) NaOH soln.

To make 500 ml of nutrient agar:

7.5 gm. of powdered agar is weighed and taken in a flask, containing 250 ml of distilled water. The content is then heated in a water-bath to allow the agar to dissolve. In another flask 1.4 gms of beef extract and 2.5 gms of peptone are dissolved in 250 ml of distilled water. The pH is then adjusted and checked with pH paper.

The two solutions are then poured in a 500 ml flask, stirred thoroughly and then heated gently. Then the medium is dispensed in culture tubes and conical flasks (as stock culture medium is autoclaved at 15 lbs. pressure for 15 min after plugging the tubes and the flask).

2. Potato-Dextrose-Agar (P.D.A.):

Potato-dextrose-agar medium is very often used for the culture of fungi. It is simply potato dextrose broth solidified by adding agar.

Freshly peeled potato – 400 gms. (40%)

(ii) Graduated cylinder – 500 ml.

(iii) Beef extract, bactopeptone, agar, and distilled water.

(iv) Balance and weight box.

(v) pH paper, comparator block 0.1(N) HCl and 0.1 (N) NaOH soln.

To make 300 ml of P.D.A.: –

4.5 gms of agar is taken in a flask containing 150 ml of distilled water. The content is heated in a water-bath to dissolve the agar. 120 gms of freshly peeled potato is taken in a flask and 150 ml water is added to it. It is boiled for 10 mins. Then this potato extract is taken and its volume is made up to 150 ml by adding water. To this extract, 7.5 gms of Dextrose is added and thor­oughly mixed.

Now the above two solutions are poured in a 500 ml flask and stirred thoroughly. This medium is dispensed in culture tubes and flasks. The tubes and flasks are plugged (Fig. 2.1.) and autoclaved.

(i) Agar must be liquefied properly before mixing with broth.

(ii) Agar must not be heated too much other­wise it will lose its ability to solidify.

(iii) Dispensing should be done quickly, otherwise the agar will solidify.

A detailed list of the composition of various media is given in the Annexure.

Step # 2. Adjustment of pH of Media:


The hydrogen-ion-concentration of culture media is of prime importance for the successful cultivation of bacteria. Some species grow best in acid medium, others in alkaline medium still others prefer substrates neutral in reaction. The H + concentration i.e.

So, for the growth of a particular macro- organism, the medium should have a specific pH. To adjust the pH, acid and alkali are used.


(ii) pH paper and colour standard

(iv) 0.1(N) NaOH and 0.1(N) HCl soln.


The simplest method for determining the pH of a soln. is to use commercially available pH paper which is impregnated with an indicator. The latter gives a colour change over a pH range of 6.4 to 8.2. A strip of pH paper is cut into small pieces and each piece is placed within a well on the comparator block.

With the help of a glass rod a drop of medium is drawn out and put on the piece of pH paper. The resulting colour is compared with the colour standard supplied.

The pH of the medium, if found to be acidic, is brought to the required pH by adding 0.1 (N) NaOH drop-wise and testing with pH paper after thoroughly mixing with a glass rod. Conversely, 0.1 (N) HCl is used to get an acidic pH of the medium.


(i) While neutralizing the broth, acid or alkali should be added drop-wise.

(ii) After adding acid or alkali the medium should be stirred to ensure proper mixing of the acid or alkali added.

(iii) At every step colour reaction should be noted.

Step # 3. Preparation of Stabs and Slants:


In order to prepare stabs, the medium is poured up to 1/2 of the culture tube (about 20 ml), which is then plugged carefully and sterilised in autoclave. After sterilization, the culture tube is kept erect in a test tube stand until the medium solidifies. Then they are collected in a wire-net basket and preserved.

In order to prepare “slants” the medium is taken up to 1/4 of a culture tube (about 7 ml) with the help of a measuring cylinder and funnel. Then the culture tubes are plugged and autoclaved. After sterilisation, before the medium sets, the tubes are sloped on a bench by leaning them against a length of a wooden stick of 1/2″ thickness in such a fashion that the medium does not touch the plug.

The culture tubes are maintained in that position until the medium solidifies. After solidification of the medium, the “slants” are collected in a wire net basket and preserved with a label indicating the date of preparation and the nature of the medium (Fig. 2.2).


(i) Culture tubes should be plugged with non-absorbent cotton.

(ii) It should be noted that during dispensing, the medium does not stick to the sides of the culture tubes.

(iii) During slanting, care should be taken to see that the medium does not touch the plug.

(iv) Stabbing or slanting should be done just before solidification i.e. at around 47°C, otherwise, there will be water inside the slant.

(v) Stabs and slants should not be disturbed before the medium solidifies.

Step # 4. Pouring of Plates:


Plate culture consists of an organism growing on a solid medium contained in a petridish. “Pouring of plates” refers to the process of pouring melted nutrient agar into petridishes. By this process a larger surface area is created for the growth of microbes in all directions.


(i) Nutrient agar “stab” or medium in flask.

(ii) Sterilised petridishes.

(iv) Absorbent cotton, rectified spirit, glass marking pencil etc.


Solid medium contained in culture tubes or flasks is melted in a water-bath. Once the agar is melted perfectly, the medium is allowed to cool around 45°C. The working table is cleaned and sterilised with cotton soaked in rectified spirit. Hands are also sterilised with rectified spirit.

The cotton plug of the melted tube or flask is opened near a flame and the mouth of the tube/flask is flamed in a semi-horizontal position. With the left hand the lid of a petridish is raised far enough to permit the mouth of the tube or flask to enter without touching the sides.

About 15 ml medium is quickly and carefully poured into the petridish, the tube/flask is withdrawn and the cover or lid is replaced. The petridish is tilted slightly with the movement of the wrist to allow homogeneous spreading of the medium (Fig. 2.3). When the medium solidifies, the plates are incubated at 30°C in an inverted position i.e. upside down so that moisture drop does not fall on the medium.


(i) All work should be done aseptically.

(ii) After solidification the plates should be kept inverted.

(iii) During pouring it should be noted that the mouth of the tube/flask does not touch any part of the petridish.

Step # 5. Inoculation of Bacteria in Nutrient Slants and Agar Plates:

Materials Required:

(i) Nutrient slants and nutrient agar plates.

(iv) Bacterial culture slant:


First the inoculating needle (Fig. 2.4) is sterilised by heating strongly in a flame and cooled by holding it outside the flame. After cooling the needle further by touching it on the ‘extra’ medium (where there is no growth) in supplied slants, a very small amount of inoculum is taken out by means of the needle and then inoculates in the previously prepared slants in a zigzag manner.

Each bacterial culture is inoculated in duplicate. The needle is then flamed to ensure killing of bacteria in the needle. (Fig. 2.5).

For streaking plates, inoculum is taken out in the same way as mentioned and touched on the solid medium in a plate. The needle is flamed to kill excess bacteria and cooled. The inoculum is spread by streaking a line with the needle (Fig. 2.6). The needle is again burnt for the same purpose and cooled.

Continuous and zigzag lines arc streaked so as to reduce gradually the number of cells along the line and to get isolated single colonies after incubation. The whole operation of inoculation is performed aseptically in front of a strong flame. The inoculated slants and plates (in inverted position) are incubated overnight at 37°C.


Bacteria are routinely cultured in a solid medium i.e. Nutrient Agar Medium (NAM) to obtain the discrete colonies of the bacteria present in the specimen or to get the information about cultural characteristics of bacteria on a solid medium, colony morphology and patterns of growth etc. The Solid basal medium, used in bacteriology laboratory constitutes the 4 essential components –

Beef Extract – It is the beef derivative which is a rich source of Organic Carbon, Nitrogen, Vitamins and Inorganic Salts that supports the rapid growth of bacterium in the laboratory at an optimum temperature, pH, and Osmotic Pressure.

Peptone – It is a semi-digested protein which is soluble in water and easily metabolized by the bacterial cell, provides the rich source of protein to the bacterial cell of the rapid growth.

Sodium Chloride – It maintains the osmotic pressure in the agar medium so that the movement of molecules takes place in and out of the bacterial cell. It must be present in right proportion otherwise it will lead to the lysis of the bacterial cell.

Agar-Agar – It is often called as Agar, is a complex polysaccharide, a carbohydrate consisting of 3, 6-Anhydro-L-galactose and D-galactopyranose, free of nitrogen, produced from various red-purple algae belonging to Gelidium, Gracilaria, Gigastina etc. It liquefies on heating to 96 °C and hardens into a jelly on cooling at 40-45 °C.

The methodology explained in this article is based on HiMedia Labs Sabouraud Dextrose Broth Medium which you can check here

The above-mentioned components can be modified by adding the various substances in a variety of ways as per the requirements of the bacterium so that the rapid and satisfactory growth of the bacterial cell takes place.

Preparing the 5X stock

Add the following reagents to a 2-liter flask:

  • 10 g (NH4)2SO4
  • 68 g KH2PO4
  • 2.5 mg FeSO4.7H2O
  • 1 liter of high quality distilled water

Once the ingredients are added, heat with stirring until the components are completely dissolved. Adjust to pH 7 with KOH. Pour the solution into smaller bottles with loosened caps and the autoclave at 15 lb/in 2 for 15 min. If you wish to add add antibiotics or nutritional supplements, do this only after the autoclave cycle is complete, as the high temperature may destroy these components. Wait until the the bottle is less than 50°C (it should be warm to touch), and then add the components. After the bottles cool to below 40°C, the caps can be tightened and the concentrated medium stored indefinitely at room temperature.

1: Media Preparation - Biology

Article Summary:

Plant Tissue Culture Media: Types, Constituents, Preparation and Selection
Author: Cornelius Onye Nichodemus


Plant tissue culture is a plant growth medium used in the laboratory for the cultivation of plant cell culture. It is usually a solid, liquid or semi-solid medium designed to support the growth of plant parts (explants), cells or microorganisms. In plant tissue culture, different types of growing media are used to cultivate various plants therefore each plant has a specific culture media best ideal for its growth and development. Plant tissue culture media (medium) is also known as growth media, culture medium, substrate etc. Generally, just like soil plant tissue culture medium plays three major roles

1. Physically support plant growth

2. Allow for maximum root growth

3. Supply root with necessities such as water, air and nutrients.

The culture media are largely responsible for the in vitro growth and morphogenesis of plant tissues. The success of the plant tissue culture depends on the choice of the nutrient medium. For the cells of most plants can be grown in a culture media. Basically, the plant tissue culture media should contain the same nutrients as required by the whole plant. It may be noted that plants in nature can synthesize their own food material. However, plants growing in vitro are mainly heterotrophic i.e. they cannot synthesize their own food.


The composition of the culture media is primarily dependent on two factors:

1. The particular species of the plant.

2. The type of material used for culture i.e. cells, tissues, organs, protoplasts.

Thus, the composition of a medium is formulated considering the specific requirements of a given culture system. The media used may be solid (solid medium) or liquid (liquid medium) in nature. The selection of solid or liquid medium is dependent on the type of plant to be grown.


There are basically five (5) types of plant tissue culture media. They are:

1. White’s medium: This is one of the earliest plant tissue culture media which was developed for root culture.

2. MS medium: Murashige and Skoog (MS) originally formulated a medium to induce organogenesis and regeneration of plants in cultured tissues. Today, MS medium is widely used for many types of culture systems.

3. B5 medium: It was developed by Gamborg, B5 medium was originally designed for cell suspension and callus cultures. At present with certain modifications, this medium is used for protoplast cultures.

4. N6 medium: Chu formulated this medium and it is used for cereal anther culture, besides other tissue cultures.

5. Nitsch’s medium: This medium was developed by Nitsch and Nitsch and frequently used for anther cultures.

Among the media explained above, MS medium is the most frequently used in plant tissue culture work due to its success with several plant species and culture systems. Interestingly, each of the above listed medium differ from each other in terms of their composition (presence or absence of some nutrients and also amount of nutrients in mg/l)

Synthetic and natural media

When a medium is composed of chemically defined components, it is referred to as a synthetic medium. On the other hand, if a medium contains chemically undefined compounds (e.g., vegetable extract, fruit juice, plant extract), it is regarded as a natural medium. Synthetic media have almost replaced the natural media for tissue culture.

Expression of concentrations in media

The concentrations of inorganic and organic constituents in culture media are usually expressed as mass values (mg/l or ppm).


Many elements are needed for plant nutrition and their physiological functions. Thus, these elements have to be supplied in the culture medium to support adequate growth of cultures in vitro. The culture media usually contain the following constituents:

2. Carbon and energy sources

The inorganic nutrients consist of macronutrients (concentration >0.5 mmol/l ) and micronutrients (concentration <0.5 mmol/l ). A wide range of mineral salts (elements) supply the macro- and micronutrients. The inorganic salts in water undergo dissociation and ionization. Consequently, one type of ion may be contributed by more than one salt. For instance, in MS medium, K ions are contributed by KNO3 and KH2PO4 while NO2 ions come from KNO3 and NH4 NO3.

Macronutrient elements: The six elements namely nitrogen, phosphorus, potassium, calcium, magnesium and sulfur are the essential macronutrients for tissue culture. The ideal concentration of nitrogen and potassium is around 25 mmol I while for calcium, phosphorus, sulfur and magnesium, it is in the range of 1-3 mmol I . For the supply of nitrogen in the medium, nitrates and ammonium salts are used together.

Micronutrients: Although their requirement is in minute quantities, micronutrients are essential for plant cells and tissues. These include iron, manganese, zinc, boron, copper and molybdenum. Among the microelements, iron requirement is very critical. Chelated forms of iron and copper are commonly used in culture media.


Plant cells and tissues in the culture medium are heterotrophic and therefore, are dependent on the external carbon for energy. Among the energy sources, sucrose is the most preferred. During the course of sterilization (by autoclaving) of the medium, sucrose gets hydrolysed to glucose and fructose. The plant cells in culture first utilize glucose and then fructose. In fact, glucose or fructose can be directly used in the culture media. It may be noted that for energy supply, glucose is as efficient as sucrose while fructose is less efficient.

It is a common observation that cultures grow better on a medium with autoclaved sucrose than on a medium with filter-sterilized sucrose. This clearly indicates that the hydrolysed products of sucrose (particularly glucose) are efficient sources of energy. Direct use of fructose in the medium subjected to autoclaving, is found to be detrimental to the growth of plant cells. Besides sucrose and glucose, other carbohydrates such as lactose, maltose, galactose, raffinose, trehalose and cellobiose have been used in culture media but with a very limited success.

The organic supplements include vitamins, amino acids, organic acids, organic extracts, activated charcoal and antibiotics.

i. Vitamins: Plant cells and tissues in culture (like the natural plants) are capable of synthesizing vitamins but in suboptimal quantities, inadequate to support growth. Therefore the medium should be supplemented with vitamins to achieve good growth of cells. The vitamins added to the media include thiamine, riboflavin, niacin, pyridoxine, folic acid, pantothenic acid, biotin, ascorbic acid, myo­inositol, Para amino benzoic acid and vitamin E.

ii. Amino acids: Although the cultured plant cells can synthesize amino acids to a certain extent, media supplemented with amino acids stimulate cell growth and help in establishment of cells lines. Further, organic nitrogen (in the form of amino acids such as L-glutamine, L-asparagine, L- arginine, L-cysteine) is more readily taken up than inorganic nitrogen by the plant cells.

iii. Organic acids: Addition of Krebs cycle intermediates such as citrate, malate, succinate or fumarate allow the growth of plant cells. Pyruvate also enhances the growth of cultured cells.

iv. Organic extracts: It has been a practice to supplement culture media with organic extracts such as yeast, casein hydrolysate, coconut milk, orange juice, tomato juice and potato extract. It is however, preferable to avoid the use of natural extracts due to high variations in the quality and quantity of growth promoting factors in them. In recent years, natural extracts have been replaced by specific organic compounds e.g., replacement of yeast extract by L-asparagine replacement of fruit extracts by L-glutamine.

v. Activated charcoal: Supplementation of the medium with activated charcoal stimulates the growth and differentiation of certain plant cells (carrot, tomato, orchids). Some toxic/inhibitory compounds (e.g. phenols) produced by cultured plants are removed (by adsorption) by activated charcoal, and this facilitates efficient cell growth in cultures. Addition of activated charcoal to certain cultures (tobacco, soybean) is found to be inhibitory, probably due to adsorption of growth stimulants such as phytohormones.

vi. Antibiotics: It is sometimes necessary to add antibiotics to the medium to prevent the growth of microorganisms. For this purpose, low concentrations of streptomycin or kanamycin are used. As far as possible, addition of antibiotics to the medium is avoided as they have an inhibitory influence oon the cell growth.

Plant hormones or phytohormones are a group of natural organic compounds that promote growth, development and differentiation of plants. Four broad classes of growth regulators or hormones are used for culture of plant cells-auxins, cytokinins, gibberellins and abscisic acid. They promote growth, differentiation and organogenesis of plant tissues in cultures.

i. Auxins: Auxins induce cell division, cell elongation, and formation of callus in cultures. At a low concentration, auxins promote root formation while at a high concentration callus formation occurs. Examples of commonly used auxins are: Indole 3-acetic acid(IAA), Indole 3-butyric acid (IBA), 1-Naphthylene acetic acid (NAA), 2, 4-Dichlorophenoxy acetic acid (2,4-D) etc. Among the auxins, 2, 4-dichlorophenoxy acetic acid is most effective and is widely used in culture media.

ii. Cytokinins: Chemically, cytokinins are derivatives of a purine namely adenine. These adenine derivatives are involved in cell division, shoot differentiation and somatic embryo formation. Cytokinins promote RNA synthesis and thus stimulate protein and enzyme activities in tissues. Examples of commonly used cytokinins are 6-Benzyl aminopurine (BAP), Benzyladenine (BA), Kinetin, Zeatin, etc. Among the cytokinins, kinetin and benzyl-amino purine are frequently used in culture media.

Ratio of auxins and cytokinins: The relative concentrations of the growth factors namely auxins and cytokinins are crucial for the morphogenesis of culture systems. When the ratio of auxins to cytokinins is high, embryogenesis, callus initiation and root initiation occur. On the other hand, for axillary and shoot proliferation, the ratio of auxins to cytokinins is low. For all practical purposes, it is considered that the formation and maintenance of callus cultures require both auxin and cytokinin, while auxin is needed for root culture and cytokinin for shoot culture. The actual concentrations of the growth regulators in culture media are variable depending on the type of tissue explant and the plant species.

iii. Gibberellins: About 20 different gibberellins have been identified as growth regulators. Of these, gibberellin A (GA3 ) is the most commonly used for tissue culture. GA promotes growth of cultured cells, enhances callus growth and induces dwarf plantlets to elongate. Gibberellins are capable of promoting or inhibiting tissue cultures, depending on the plant species. They usually inhibit adventitious root and shoot formation.

iv. Abscisic acid (ABA): The callus growth of cultures may be stimulated or inhibited by ABA. This largely depends on the nature of the plant species. Abscisic acid is an important growth regulation for induction of embryogenesis.

For the preparation of semi-solid or solid tissue culture media, solidifying or gelling agents are required. In fact, solidifying agents extend support to tissues growing in the static conditions.

Agar: Agar, a polysaccharide obtained from seaweeds, is most commonly used as a gelling agent for the following reasons.

1. It does not react with media constituents.

2. It is not digested by plant enzymes and is stable at culture temperature.

Agar at a concentration of 0.5 to 1% in the medium can form a gel.

Gelatin: It is used at a high concentration (10%) with a limited success. This is mainly because gelatin melts at low temperature (25°C), and consequently the gelling property is lost.

Other gelling agents: Bio-gel (polyacrylamide pellets), phytagel, gelrite and purified agarose are other solidifying agents, although less frequently used. It is in fact advantageous to use synthetic gelling compounds, since they can form gels at a relatively low concentration (1.0 to 2.5 g l ).

The optimal pH for most tissue cultures is in the range of 5.0-6.0. The pH generally falls by 0.3-0.5 units after autoclaving. Before sterilization, pH can be adjusted to the required optimal level while preparing the medium. It is usually not necessary to use buffers for the pH maintenance of culture media.

At a pH higher than 7.0 and lower than 4.5, the plant cells stop growing in cultures. If the pH falls during the plant tissue culture, then fresh medium should be prepared. In general, pH above 6.0 gives the medium hard appearance, while pH below 5.0 does not allow gelling of the medium.


The general methodology for a medium preparation involves preparation of stock solutions (in the range of 10x to 10Ox concentrations) using high purity chemicals and demineralized water. The stock solutions can be stored (in glass or plastic containers) frozen and used as and when required. Most of the growth regulators are not soluble in water. They have to be dissolved in NaOH or alcohol.

Dry powders in Media Preparation: The conventional procedure for media preparation is tedious and time consuming. Now a days, plant tissue culture media are commercially prepared, and are available in the market as dry powders. The requisite medium can be prepared by dissolving the powder in a glass distilled or demineralized water. Sugar, organic supplements and agar (melted) are added, pH adjusted and the medium diluted to a final volume (usually 1 litre).

Sterilization of Media: The culture medium is usually sterilized in an autoclave at 121°C and 15 psi for 20 minutes. Hormones and other heat sensitive organic compounds are filter-sterilized, and added to the autoclaved medium.


In order to select a suitable medium for a particular plant culture system, it is customary to start with a known medium (e.g. MS medium, B5 medium) and then develop a new medium with the desired characteristics. Among the constituents of a medium, growth regulators (auxins, cytokinins) are highly variable depending on the culture system.

In practice, 3-5 different concentrations of growth regulators in different combinations are used and the best among them are selected. For the selection of appropriate concentrations of minerals and organic constituents in the medium, similar approach referred above, can be employed.

Medium-utmost Important for Culture: For tissue culture techniques, it is absolutely essential that the medium preparation and composition are carefully followed. Any mistake in the preparation of the medium is likely to do a great harm to the culture system as a whole.

About Author / Additional Info:
I am a First Class graduate of Plant Science and Biotechnology from University of port Harcourt.

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Research Benefits from an Online Presence

In the age of the internet, social media tools offer a powerful way for scientists to boost their professional profile and act as a public voice for science. Although the type of online conversations and shared content can vary widely, scientists are increasingly using social media as a way to share journal articles, advertise their thoughts and scientific opinions, post updates from conferences and meetings, and circulate information about professional opportunities and upcoming events. Google searches now represent the standard approach for discovering information about a topic or person—whether it be search committees collecting information about faculty candidates, graduate students searching out prospective labs, or journalists on the hunt for an expert source. Consequently, in today's technology-driven world, lack of an online presence can severely limit a researcher's visibility, and runs the risk that undesirable search results appear before desirable ones (however, this scenario is easily rectified see Box 2). A growing body of evidence suggests that public visibility and constructive conversation on social media networks can be beneficial for scientists, impacting research in a number of key ways.

Box 2. Advice for New Users

In academia, there is often a particular stigma attached to online activities. Actively maintaining an online profile and participating in social media discussions can be seen as a waste of time and a distraction from research and teaching duties. We believe this perception is misguided and based on incorrect interpretations of what scientists are actually doing online. When used in a targeted and streamlined manner, social media tools can complement and enhance a researcher's career. When exploring online tools for the first time, new users can maximize their reach by considering the following points:

Practical Work for Learning

The ability to make different culture media for culturing different microorganisms is an essential part of any microbiology investigation.

Agar provides a matrix of supporting jelly for dissolved nutrients. Different nutrients are appropriate for culturing different microorganisms.

Apparatus and Chemicals

Add different nutrients to the basic agar depending on the organisms you are planning to culture.

Health & Safety and Technical notes

Refer to CLEAPSS Recipe card 1 for full details of handling agar. The identified hazard relates to inhaling the powder when making up the agar jelly – the control measure is weighing out the powder in a fume cupboard. The risk of scalds when dealing with hot liquid is reduced by wearing oven gloves.

1 Basic agar Mix 1.5 g of agar with 10 cm 3 of water into a paste. Slowly add more water with stirring until the volume is 100 cm 3 . Heat the mixture on a boiling water bath to 95 °C in the required container. This preliminary heating can be omitted if the agar will then be sterilised, unless it is necessary to decant the agar into smaller containers prior to autoclaving.

2 Nutrient agar for bacteria Mix 2 g of Bovril, 0.5 g of sodium chloride, and 1.5 g of agar with 10 cm 3 of water into a paste. Slowly add more water while stirring until the volume is 100 cm 3 . Heat in a boiling water bath to 95 °C in the required container.

3 Malt agar for fungi Mix 2 g of malt extract with 2 g of agar with 10 cm 3 of water into a paste. Slowly add more water with stirring until the volume is 100 cm 3 . Heat in a boiling water bath to 95 °C in the required container.

4 Starch agar Make a paste containing 1 g of soluble starch in 10 cm 3 of hot water. Add 1.5 g of agar, stir well, and slowly add more water with stirring until the volume is 100 cm 3 . Heat in a boiling water bath to 95 °C in the required container.

5 Starch malt agar for growth of fungi and digestion of starch Mix 3 g of light malt (crystal or spraymalt) powder (from home-brewing shops), and 0.5 g of peptone (to promote growth) in 20 ml of water. Also make a paste containing 1 g of soluble starch in 10 cm 3 of hot water. Add these two solutions to 1.5 g of agar while stirring, and slowly add more water with stirring until the volume is 100 cm 3 . Stir before decanting into smaller containers (if required) and sterilising.

6 Mannitol yeast extract agar (MYEA) For 1 litre of mannitol yeast extract agar (MYEA) suspend 10 g of agar in 1 litre of water. Heat to dissolve. Add 0.5 g K2HPO4 (Hazcard 95C), 0.2 g MgSO4.7 H2O (Hazcard 59B), 0.2 g NaCl (Hazcard 47B), 0.2 g CaCl2.6H2O (Hazcard 19A), 10 g mannitol, and 0.4 g yeast extract. K2HPO4, MgSO4.7 H2O and NaCl are described as low hazard. CaCl2.6H2O is an irritant as a powder, but not in the final medium.

7 Nitrogen-free mineral salts agar For 500 cm 3 , first dissolve 0.05 g FeCl3.6H2O in 500 cm 3 distilled water. Add 2 g K2HPO4, 0.25 g MgSO4.7H2O, and 10 g glucose. Dissolve and check pH, adjusting to 8.3 if necessary with 0.1M NaOH (Refer to CLEAPSS Hazcard 91: Irritant at this concentration). Pour into a bottle containing 1 g CaCO3 and 7.5 g agar. Autoclave at 121°C. Mix to disperse the CaCO3 before pouring. You can buy ready-made nitrogen-free mineral salts agar. K2HPO4 (Hazcard 95C), MgSO4.7H2O (Hazcard 59B), glucose (Hazcard 40C), and CaCO3 (Hazcard 19B) are all listed as Low hazard. FeCl3.6H2O is harmful as a solid, but not at the low concentration of the final agar.


CLEAPSS Laboratory Handbook section 15.2.7 has more information on how to manage agar and prepare plates. Or you can buy prepared media in bottles or plates (see Suppliers below).

a Calculate the quantity required and prepare just enough agar for the investigation – around 15 cm 3 for normal depth in a 90 mm Petri dish. Any surplus will keep for 6-12 months in tightly-sealed screw-top bottles if sterile.

b Weigh out the agar medium powder containing the gel and chosen nutrients, add water and sterilise the mixture for the time, and at the temperature, specified by the manufacturer.

c Heat agar and water at 95 °C to dissolve the agar. Always use a water bath to boil agar, and never add agar to boiling water.

d Stopper flasks with a well-fitting plug of non-absorbent cotton wool. Cover with greaseproof paper or aluminium foil before sterilising by autoclaving.

e After autoclaving, transfer to a water bath to equilibrate at 50 °C. Stack plates after pouring to minimise condensation except in the top plate(s).

f Warm the Petri dishes before pouring to minimise condensation.

g Keep poured plates in a sealed plastic bag until needed to reduce dehydration of the media.

h Divide the agar into individual sterile McCartney bottles if you want the students to pour their own plates (see Pouring an agar plate).

Web links

Microbiology teacher resources
Society for General Microbiology – source of Basic Practical Microbiology, an excellent manual of laboratory techniques and Practical Microbiology for Secondary Schools, a selection of tried and tested practicals using microorganisms.

Microbiology online
MiSAC (Microbiology in Schools Advisory Committee) is supported by the Society for General Microbiology (see above) and their websites include more safety information and a link to ask for advice by email.

(Websites accessed October 2011)

© 2019, Royal Society of Biology, 1 Naoroji Street, London WC1X 0GB Registered Charity No. 277981, Incorporated by Royal Charter

How to Make a Cell Lysis Solution

The precise components and procedure required for making a cell lysis solution depends on several factors, including the type of cells and the objective of the experiment. This BiologyWise article explains how to make cell lysis solution with respect to the major ingredients, and how they vary with different experimental setups.

The precise components and procedure required for making a cell lysis solution depends on several factors, including the type of cells and the objective of the experiment. This BiologyWise article explains how to make cell lysis solution with respect to the major ingredients, and how they vary with different experimental setups.

Sodium dodecyl sulfate (SDS), the most commonly used detergent for cell lysis and protein denaturation, is also widely used in several soaps, shampoos, laundry detergents, and toothpastes.

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A cell lysis solution is a detergent-based buffer solution used to break open the desired cells and further isolate a particular cellular component of interest. It is also referred to as a cell lysis buffer or simply, lysis buffer. This process of lysing cells using chemical agents is termed as chemical disruption.

Apart from detergent and buffering agents, additional agents are added that aid the lysing process, eliminate unwanted cellular components, and/or protect the desired cellular component. These additions depend on the cell type involved, the cellular component to be studied, and the precise techniques that need to be performed on the lysate.

Given below is the basic procedure to prepare a cell lysis solution, followed by a description of the components required to prepare the solutions, and their variations with respect to the commonly followed protocols for different experiments.

Procedure to Make a Cell Lysis Solution

Given below is the procedure to prepare a lysis solution containing 10mM Tris-HCl buffer, 1mM EDTA as the chelating agent, and 0.5% SDS as the detergent.

Dissolve 121 g Tris-HCL (molecular weight = 157.60 g) in 800 ml distilled water, adjust the pH to 8 using HCl solution, and make up the volume to 1 L using distilled water.

Step 2: Preparation of 500 ml of 0.5 M EDTA stock solution

Dissolve 93.0 g of EDTA [EDTA.Na2.2H2O] (molecular weight = 372.24 g)] in 400 ml of distilled water, add 10 g (approx.) NaOH pellet to adjust pH to 8, and make up the volume to 500 ml using distilled water.

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Step 3: Preparation of 10% SDS stock solution

Dissolve 10 g of SDS in 90 ml distilled water, and make up the volume to 100 ml using distilled water.

Step 4: Preparation of 500 ml of the Tris-EDTA SDS lysis buffer

Choosing the Reagents for a Cell Lysis Buffer

All cell lysis solutions are prepared using a suitable buffer solution, so as to maintain the appropriate pH. Disruption of the cells will disturb the internal pH of the cell, which alters the structural integrity of proteins and other macromolecules. In order to avoid this, generally a buffer solution adjusted at a pH equal to the normal intracellular pH is used.

A buffer solution comprises a weak base and its conjugate acid, or a weak acid and its conjugate base. Each buffering agent can work as an effective buffer only within a specific pH range called the buffer range. The choice of buffering agent depends on this buffer range. Generally, the buffers are used at a concentration of 10 – 50 mM and are adjusted to pH 7.4 – 8.

The two commonly used buffers for making cell lysis solutions, and their buffer range are as follows:

Cell membranes are made up of phospholipid layers which are held together through polar interactions. A detergent serves as an emulsifier and disrupts these polar interactions. It also forms complexes with the lipid molecules and protein molecules embedded in the membrane, and precipitates them out of the solution.

Detergents can be grouped as ionic or non-ionic, depending on the nature of the hydrophilic group, or as mild or strong depending on their ability to solubilize membranes and proteins. Ionic detergents with positively charged groups are termed cationic detergents, and those with negatively charged groups are termed anionic detergents.

Mild detergents are used when cells need to be lysed for purifying and/or studying cellular proteins, so as to avoid denaturation of the desired proteins. On the other hand, strong detergents that denature cellular proteins are used while lysing cells for DNA isolation.

Given below are some of the commonly used detergents, their application as cell lysing agents, and usual concentrations in which they are used for cell lysis.

1) SDS (sodium dodecyl sulfate) [anionic]

General use: DNA isolation from any cell type, protein isolation under denaturing conditions
Concentration: 1 – 10 %

2) Sodium Deoxycholate [anionic]

General use: Purification of membrane proteins
Concentration: 0.5 %

3) CTAB (cetyltrimethylammonium bromide) [cationic]

1) NP-40 (nonyl phenoxypolyethoxylethanol)

General use: Nuclei isolation
Concentration: 0.1 – 1.0 %

2) Triton X-100

General use: Purification of membrane proteins, DNA isolation
Concentration: 0.1 – 5.0 %

3) Polysorbate 20

General use: Purification of membrane proteins, immunoprecipitation
Concentration: 0.05 – 0.5 %

Role: The chelating agent is a chemical that sequesters divalent cations, like Mg ++ , and Ca ++ , which are required for membrane stability. Due to such chelation, these cations are no longer associated with the membrane, thereby, weakening it. Moreover, the lysing of membrane results in exposing the nucleic acids to nucleases, which are otherwise sequestered inside the lysosomes. These nucleases are inactive in the absence of divalent ions. Thus, chelating agents protect the DNA and RNA molecules from degradation by these nucleases.

Osmotic Stabilizers: Glucose, sucrose, sorbitol, and glycerol may be added for osmotic stability, and preventing the cellular components from osmotic shock that may occur during the sudden rupturing of cell membrane. In addition, they also help stabilize lysosomal membranes, thereby, reducing the release of degradative enzymes from the lysosomal lumen.

Salts: Disruption of cells and the release of cellular components into the medium, may alter the ionic strength of the medium. In order to deal with this and maintain ionic strength, salts, like sodium chloride [NaCl], potassium chloride [KCl], and ammonium sulfate [(NH4)2SO4] may be added.

Enzymes: Depending on the target macromolecules to be isolated from cells, lytic enzymes may be added to either protect them from the action of other enzymes, or degrade the remaining macromolecules that may contaminate the lysate.

Cell lysis solutions are used for the chemical disruption of cells, and may also be used in a combination with other methods, like homogenization, sonication, grinding, freeze-thaw techniques, etc. This is especially useful for lysis of fungal cells and plant cells, since they have strong cell walls around the cell membranes.

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How do you guys prepare 10% FBS Medium? - (Nov/27/2008 )

The people in my lab just add 50ml of FBS and 5 ml Pen Strep in 500ml DMEM. They say that the difference in cell growth is minimal. But i was taught to keep using the same lot of FBS throughout my whole experiment. But if the effect of different concentration of FBS on cell growth is minimal, what is the point of keep using the same lot of serum?

The people in my lab just add 50ml of FBS and 5 ml Pen Strep in 500ml DMEM. They say that the difference in cell growth is minimal. But i was taught to keep using the same lot of FBS throughout my whole experiment. But if the effect of different concentration of FBS on cell growth is minimal, what is the point of keep using the same lot of serum?

If a bottle of DMEM/RPMI is 500ml and you need to make up a 10% FBS/FCS solution . then you need to add 55mls of FBS/FCS to the 500ml of media. If you do not then you are adding a 10% error into your experiment straight away.

You need to batch test serum. this is extremely important as there is MASSIVE batch variation . and MASSIVE differences in FBS/FCS origins.

NZ>Austalian>North American>European>South American>British

If you look at Vaibility and growth then there will be no differences between the above. HOWEVER these are insensitive methods for looking at the "health of the cells". If you look at other cell markers, then you will see massive variations:

Oxygen Consumption
Enzyme Induction
Receptor Expression
Cytochrome C contant
Electron transport chain complexes

Rhombus, that is a damn fine answer, especially as I am a Kiwi!

I am glad you like the answer. NZ serum has always been the best in the world. I do not know why. However experimentally it is vital for us to have good cells. For an example, we have HEK cells Sodium channel expression. with NZ serum as the only varaible, we find 60% greater expression in cells grown in NZ serum compared to other sources of serum. IMPORTANT when you are doing patch clamp work

Watch the video: Media Preparation Session 12 - Alliance Bio Expertise (January 2022).