Determination of BOD of water

Determination of BOD of water

Aim

To determine the dissolved oxygen concentration and biological oxygen demand of pond water

Principle

Dissolved oxygen of water is of paramount importance to all living organisms and is considered to be a factor which to a greater extent can reveal the nature of the whole aquatic system at a glance.  Dissolved oxygen is used as an indicator of the health of a water body.  Higher dissolved oxygen concentrations are correlated with high productivity and little pollution.  The presence of DO in water is mainly due to Direct diffusion from air and by Photosynthetic evolution by aquatic autotrophs.

The first one is purely a physical process and depends on the solubility of oxygen under the influence of temperature, salinity, water movement, whereas the later is a biological process and depends on the availability of light and rate of metabolic processes resulting in diurnal fluctuations.

Two methods are commonly used to determine DO concentration: The iodometric method which is a titration-based method and The membrane electrode procedure.  In the Iodometric method, divalent manganese solution is added to water, followed by addition of strong alkali in a glass-stopper bottle. DO rapidly oxidize an equivalent amount of the dispersed divalent manganese hydroxide precipitates to hydroxides of higher valence states. In acidic conditions, in the presence of iodide ions, the oxidized manganese reverts to the divalent state and iodine equivalent of the original DO content get liberated. The iodine is then titrated with a stranded solution of thiosulfate and the titration end point can be detected using a starch indicator.  Winkler Method uses titration to determine dissolved oxygen in the water sample. 

Biochemical oxygen demand (BOD) is the amount of dissolved oxygen needed or demanded by aerobic organisms to break down organic material present in a given sample at certain temperature over a specific time period.  It is a good index of organic pollution.  If the amount of organic matter in a sewage is more, then more oxygen will be utilised by bacteria to degrade it.  

BOD is calculated by keeping a sample of water containing a known amount of dissolved oxygen for five days in dark under aerobic conditions at 20^OC.   The oxygen content is measured again and BOD calculated.  Since nitrification consumes oxygen significantly, it will result in over estimation of BOD and must be checked by adding 1 ml of 0.5% solution of Allyl thourea.  In the water Samples where more than 70% of initial oxygen is consumed, it is necessary to aerate/oxygenate/dilute the sample with BOD free water to avoid oxygen stress.

BOD level in mg/litre                        Quality of water

            1 - 2                                         - Very Good

            3 - 5                                         - Fair

            6-9                                           - Poor

            100 or more                             - Very Poor

Reagents required

·         Manganese Sulphate solution - 48 g MnSO4.4H2O or 40 g MnSO4.2H2O or 36,4 g MnSO4.H2O in 100 ml distilled water

·         Alkali - iodide – azide reagent

o   Solution A - 500 mg NaOH and 135 g Sosium Iodide in 1 litre distilled water

o   Solution B - 10 g Sodium Azide in 40 ml distilled water

o   Solution A and B are mixed to get alkali -iodide - azide solution

·         Concentrated sulphuric acid

·         0.025 N Sodium Thiosulphate solution

·         Starch Solution - 2 g soluble starch in 100 ml distilled water.

·         Allyl Thio Urea Solution (0.05%)

·         Sulphuric acid (1N) - 2.8 ml of concentrated H2SO4 added to 100 ml distilled water

·         Sodium hydroxide (1 N) - 4 g of NaOH in 100 ml distilled water

Materials required

Water Sample, Pipette, Titration assembly, BOD bottle, BOD Incubator, Flasks, etc

Procedure

• pH of water sample was adjusted to neutral using 1N acid or 1 N alkali Solution.
• The pond water sample was filled in 2 BOD bottles without any bubble formation.
• 1 ml alkyl thiourea was added to 1 bottle.
• 2 ml of manganese sulphate was added slowly to the bottle by inserting pipette below the surface of water so that no air bubbles are introduced via the pipette.
• 2 ml of alkali - iodide - azide reagent was added in the same manner.  The bottle was stoppered with care so as not to allow air entry in to the bottle and mixed the sample by inverting several times.  If oxygen is present, a brownish - orange colour of precipitate or floc will appear.  
• The precipitates formed are dissolved by adding 2 ml of concentrated sulphuric acid by keeping the pipette very near to sample surface.  Again after carefully stoppering, the bottle was inverted several times to dissolve the floc.  
• 50 ml of the sample from the bottle was titrated with 0.025 N sodium thiosulphate, till a pale straw colour is formed.
• 2 ml of starch solution was added, and a blue colour formed
• Titration is continued until the sample colour turns clear.
• The burette reading was noted.

·         Calculation of D.O. in mg/Liter = 8 X 100 X N/V X v

§  V = Volume of sample taken 

§  v = Volume of 0.025N Sodium thiosulphate solution used 

§  N = Normality of titrant = 0.025

§  8 is constant (1ml of 0.025N Sodium thiosulphate solution is equivalent to 0.2mg oxygen).

• The concentration of dissolved oxygen in the sample is equivalnet to the volume of 0.025 N sodium thiosulphate used.   Each ml of 0.025 N sodium thiosulphate added is equal to 1 mg/L Dissolved oxygen. This is recorded as Dissolved Oxygen (D1).  
• The other 2 bottles are kept in BOD incubator at 20^OC and after 5 days, the dissolved oxygen was determined by the above method and the value was recorded as Dissolved Oxygen (D2).
• The BOD of water in mg/Liter was calculated as BOD = D1-D2
• D1 is the initial dissolved oxygen concentration of the sample before 5 days incubation and D2 is the concentration after 5 days of incubation.

 

 

 

 

 

Isolation of Rhizobium

 Isolation of Rhizobium 

Aim

To isolate Rhizobium species from root nodules of legumes

Principle

Nitrogen fixation is the phase of the nitrogen cycle during which enzymatically competent microorganisms (with nitrogenase) convert atmospheric nitrogen into nitrogenous compounds. This process replenishes the soil with usable nitrogen that is rapidly removed by plant use, denitrification and leaching. Nitrogen fixation is mediated by two microbial systems. One consists of non-symbiotic free living microorganisms such as members of Azotobacter, Beijerinckia, Clostridium and Cyanobacteria, which are capable of using Nitrogen gas as their nitrogen source. The second system involve symbiotic microbial forms such as members of Rhizobium, which following infection grow in a nodule in the root of leguminous plants. A mutually benefited association is established in which nutrients in the plant sap is used by Rhizobium as it fixes atmospheric nitrogen to ammonia which can be assimilated by the plants into various proteins. Rhizobium finds application as biofertilizer in recent times.

Rhizobium species are generally cultured aerobically at 25-30⁰C in Yeast extract mannitol agar (YEMA) medium. Fast growing Rhizobia develop colonies in 3-6 days and slow growing Rhizobia in 7-10 days. Congo Red is used as a dye to enable differentiation of Rhizobium species from other bacteria. Rhizobium appear as pale coloured colonies while soil bacteria such as Agrobacterium species and others absorb the red dye. YEMA is widely used for the cultivation of Agrobacterium, Rhizobium and other soil microorganisms. It contains mannitol as a carbon source and yeast extract as a source of readily available amino acids, vitamin B complex and growth factors for Rhizobia. Congo red. Colonies of Rhizobia stand out as colourless, translucent, glistening and elevated, with entire margins

Rhizobium can fix atmospheric nitrogen only in root nodules of legumes where it is in the bacteroid stage of its life cycle. It possesses the entire complement of genes for nitrogen fixation, which are normally latent and become active only under special conditions. Rhizobium makes nitrogen available to the plant and in turn, the bacteria derive nutrients from the tissues of the plants. A crushed leguminous root nodule may be used as the source of a stained slide preparation. Rhizobium appear as bacteroids within the nodule, and are distinguished by their pleomorphism, wherein they appear in a variety of shapes, such as x, y, t, v etc.

Materials Required

Freshly picked leguminous plants with root nodules as source of Rhizobium, Yeast extract mannitol agar (YEMA) plates (pH 6.8-7), 0.1% acidified mercuric chloride, 95% ethanol, methylene blue, crystal violet, safranin, Gram’s Iodine

Equipments – Bunsen burner, inoculating loop, glass slide, staining tray, glassware marking pencil, blade, glass rod.

Procedure

1.      Healthy root nodules of a young leguminous plants were obtained by cutting with a blade

2.      The nodules were washed thoroughly first with tap water, then with sterile distilled water, under aseptic conditions so as to remove contaminants and adhering soil particles

3.      Thereafter the nodules were immersed into 0.1% acidified mercuric chloride for 5 minutes

4.      The nodules were then transferred into a sterile beaker containing 10 ml of 95% ethanol and kept for 2-3 minutes and blot dried

5.      The nodules were finally washed for 5 minutes with sterile tap water using sterile blotting paper

6.      Nodules were aseptically crushed with glass rod

7.      The nodules were streaked on the YEMA medium incorporated with Congo Red

8.      Plates were incubated in an inverted position at 28⁰C for 48-72 hours

Microscopic Observation

1.      A root nodule was thoroughly rinsed and crushed between two slides.

2.      A loopful of the material from the nodule was used for preparing smears on 2 slides

3.      Gram staining was done

4.      The slides were observed under the microscope

Observation and Result

            On YEMA plates, growth was obtained on 2-3 days of the incubation.  Rhizobium appeared as gummy cream coloured colonies as it does not absorb Congo red dye. Agrobacteria and other soil bacteria appeared red since they take up the dye strongly.

On microscopic observation, Rhizobia appeared as Gram negative, non-motile, rod shaped bacteria

 

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Isolation of Azotobacter

 Isolation of Azotobacter 

Aim

To isolate Azotobacter species from soil

Principle

The free living bacteria having the ability to fix molecular nitrogen can be distinguished into obligate aerobic and facultative anaerobic organisms. Members of the genus Azotobacter of the family Azotobacteriaceae, are heterotrophic obligate aerobic nitogen fixers. Azotobacter species are Gram- negative, aerobic soil-dwelling bacteria. They are grouped into three genera namely Azotobacter, Beijerinckia and Derxia. Several species of Azotobacter are recognized in neutral and alkaline soils, in water, and in the rhizosphere of some plants. Azotobacter chroococcum, Azotobacter agilis, Azotobacter vinelandii are some common examples.

Azotobacter species are large, usually oval, but may appear in different shapes from rods to spheres, Gram negative. The size of the cells ranges from 2-10 µm long and 1-2 µm wide. They are typically polymorphic, i.e. of different sizes and shapes. In microscopic preparations, the cells can be dispersed or in irregular clusters or chains of varying lengths. In fresh cultures, cells are motile due to peritrichous flagella. Later, the cells lose their mobility, become almost spherical, and produce a thick layer of mucus, forming the cell capsule. Azotobacter are free-living in soil and water. The nitrogen fixing enzyme, nitrogenase is oxygen-sensitive. The high respiration rate of Azotobacter uses up free oxygen within the cells and protects the nitrogenase. Azotobacter species are relatively easy to isolate from soil by growing on nitrogen free media, where the bacteria are forced to use atmospheric nitrogen gas for cellular protein synthesis. Cell proteins are mineralized in soil after the death of the Azotobacter cells, enhancing the nitrogen availability of the crop plants.

Azotobacter species can be used as an important biofertilizer. Inoculation of soil or seeding with Azotobacter is effective in increasing crop yield. It is also known to synthesize biologically active substances such as vitamins, IAA, and Gibberellins in pure culture. Fungistatic properties against certain pathogenic fungi such as Alternaria, Fusarium species etc. is also reported. These attributes explain the observed effects of Azotobacter in improving seed germination and plant growth.

Materials Required

Soil sample, Jensen’s medium, 10 ml and 9 ml sterile water blanks, Sterile 1 ml pippete, sterile petridishes, Bunsen burner, glass marking pencil

Procedure

1. Jensen’s medium was prepared and sterilized by autoclaving at 121⁰C 15 lbs per inch² pressure for 15 min.

2. 1 gm of sieved soil was added to 10 ml sterile water blanks and shaken for 15-20 min.

3. Serial dilution was performed to obtain 10^-2, 10^-3. 10^-4, 10^-5 dilutions.

4. 1 ml aliquots of various dilutions were pipetted onto sterile petriplates and pour plated by adding molten medium at bearable warmth.

5. The plates were rotated for uniform distribution and incubated at 28⁰C for 3 days.

Observation

The plates were observed after 3 days incubation for the appearance of colonies on the agar surface.

Result

On Jensen’s medium, Azotobacter colonies appeared as milky and mucoid. On microscopic observation, short rods of Azotobacter were confirmed.

Demonstration of bacterial pigmentation

 

Demonstration of bacterial pigmentation

Aim:

To observe pigmentation by various bacteria when grown on nutrient agar.

Principle:

When a single bacterial cell is placed on a solid or liquid medium, it begins to divide and eventually, a colony appears in the place. Microorganisms exhibit visible physical differences in appearance in their colonies such as colonial shape and appearance, pigmentation and smell and their growth pattern when grown on different types of media. These differences are called cultural characteristics or morphology and may be used as an aid in identifying and classifying them. Cultural characteristics or morphology are determined by observing microorganisms cultured in nutrient broth and on nutrient agar plates, after incubation.

Chromogenic bacteria produce intracellular pigments that are responsible for the color of the colonies on the agar surface. Some bacteria produce extracellular soluble pigments that are excreted into the medium and that also produce a colored colony. Most microorganisms are nonchromogenic and will appear cream, white, or gray.  Pigmented bacteria are also known as chromobacteria. Bacterial pigments are water soluble or insoluble; water soluble pigments are diffused in the growth medium. Chemically, bacterial pigments are pyrrole, phenazine, carotenoid, xanthophylls and quinine or quinone derivatives. The pigment molecules are synthesized in cell wall or periplasmic space. Only aerobic and facultatively aerobic bacteria are pigmented because, molecular oxygen is essential for pigmentation. Pigment synthesis is also dependent on light, pH, temperature and media constituents like indicator dyes.  In bacteria, pigment formation is associated with morphological characteristics, cellular activities, pathogenesis, protection and survival.

Pigment                                              Bacteria

Purple                                                  Spirillum rubrum

Violet                                                  Chromobacterium violacein

Yellow                                                Xanthomonas campestris

Orange                                                Sarcina aurentiaca

Red                                                     Serratia marcescens

Black                                                   Prevotela melaninogenica

Golden                                                Staphylococcus aureus

Pink                                                     Micrococcus roseus

Fluorescent blue/green                        Pseudomonas aeruginosa

Fluorescent yellow                              Pseudomonas fluorescens

S. aureus is a facultatively anaerobic, Gram-positive coccus, which appears as grape-like clusters when viewed through a microscope, and has round, usually golden-yellow colonies, often with hemolysis, when grown on blood agar plates. The golden appearance is the etymological root of the bacterium's name; aureus means "golden" in Latin.   Some strains of Staphylococcus aureus are capable of producing staphyloxanthin - a golden coloured carotenoid pigment. This pigment acts as a virulence factor, primarily by being a bacterial antioxidant which helps the microbe evade the reactive oxygen species which the host immune system uses to kill pathogens.

Serratia marcescens is a motile,short rod-shaped, Gram-negative, facultative anaerobe bacterium.  S. marcescens produces a reddish-orange tripyrrole pigment called prodigiosin.  Prodigiosin is made up of three pyrrole rings and is not produced at 37°C, but at temperatures below 30°C.

Pseudomonas aeruginosa is a common gram-negative, rod-shaped bacterium.  P. aeruginosa can secrete a variety of pigments, including pyocyanin (blue-green), pyoverdine (yellow-green andfluorescent), and pyorubin (red-brown). The species name aeruginosa is a Latin word meaning verdigris ("copper rust"), referring to the blue-green color of laboratory cultures of the species. This blue-green pigment is a combination of two metabolites of P. aeruginosa, pyocyanin (blue) and pyoverdine (green), which impart the blue-green characteristic color of cultures.

Materials required:

Routine microbiological facilities

Nutrient agar plates

Overnight cultures of Pseudomonas aeruginosa, Serratia marcescens, Staphylococcus aureus

Procedure:

Using an inoculation loop nutrient agar plates were inoculated with Pseudomonas aeruginosa, Serratia marcescens and Staphylococcus aureus.  Two plates were kept for Serratia marcescens culture.

All plates were kept at 37oC for overnight incubation.  One set of Serratia inoculated plate were kept overnight at room temperature.

After incubation, the plates were observed for colony morphology.

Observation:

S. aureus formed medium sized, round, golden yellow pigmented colonies

Pseudomonas aeruginosa colonies were large, translucent, with a diffused greenish blue pigment.

Serratia marcescens formed medium sized, round, bright red pigmented colonies

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Bacterial Growth curve - Lab experiment

 

Bacterial Growth curve

Aim

To plot the growth curve of an aerated bacterial culture and determine the generation time

Principle

Bacterial population growth studies require inoculation of viable cells into a sterile broth medium and incubation of the culture under optimum temperature, pH and gaseous conditions. Under these conditions, the cells will reproduce rapidly. The increase in the cell number versus the time of incubation is plotted to obtain the growth curve.

            The curve can be used to explain the stages of the growth cycle. It also helps in the measurement of cell numbers and the rate of growth of a particular organism under standard conditions. The growth rate is expressed by its generation time, the time required for the microbial population to double.

            The stages of a typical growth curve are:

1.      LAG PHASE

During this stage, cells are adjusting to their new environment. Cellular metabolism is accelerated resulting in rapid biosynthesis or preparation for the next phase of the cycle. Although the cells are increasing in size, there is no cell division and therefore no increase in number.

2.      LOGARITHMIC PHASE

Under optimal nutritional and physical conditions, the physiologically robust cells reproduce at a uniform and rapid rate by binary fission. Thus there is a rapid exponential increase in the population which doubles regularly until a maximum number of cells is reached. The time required for the population to double is the generation time. The length of the log phase varies depending on the organisms and the composition of the medium. The average may be estimated to last for 6-12 hours.

3.      STATIONERY PHASE

During this stage some cells are dying. Therefore, there is no further increase in cell numbers and the population is maintained at its maximum level for a period of time. The primary factors responsible for this stage are the depletion of some essential; metabolites and accumulation of toxic, acidic or alkaline end products in the medium.

4.      DECLINE /DEATH PHASE

Because of the continuing depletion of nutrients and buildup of metabolic wastes, the microorganism die at a rapid and uniform rate. The decrease in population closely parallel its increase during log phase. Theoretically, the entire population should die during a time interval equal to that of the log phase. This does not occur because a small number of highly resistant organisms persist for an indeterminate length of time.

      Construction of a completed bacterial growth curve requires that aliquots of a 24-hour shake-flask culture be measured for population size at intervals during the incubation period. Such a procedure does not tend itself to a regular laboratory session. Therefore, this experiment follows a modified procedure designed to demonstrate only log and lag phases, the curve will be plotted by using the two values for the measurement of growth. The direct method requires enumeration of viable cells in serially diluted samples of the test culture taken at 30 minutes interval.

The indirect method uses spectrophotometric measurement of the developed turbidity at the same 30 minutes as an index of increasing cellular mass. Indirect determination of generation time is made by simple extrapolation from the log phase as illustrated in the figure. Select two points on the optical density scale such as 0.2 and 0.4 that represent doubling of the turbidity. Using a regular scale extrapolate, draw a line between each of the selected optical density on the coordinates of the plotted line of the growth curve. Then draw perpendicular lines from these end points on the Y axis to their respective time intervals on the X axis. Generation time is calculated as,

GT = t _OD0.4- t _OD0.2

Materials Required

1.      Culture- 10-12 hours (log phase) nutrient broth culture of Escherichia coli

2.      Media- 100 ml nutrient broth in 250 ml flask

3.      Equipments- 37^oC water bath shaker, incubator, spectrophotometer, Bunsen burner, glassware marking pencil

Procedure

1.      Nutrient broth was prepared and 100 ml medium dispersed into series of 250 ml conical flask and sterilized at 121^oC and 15 lbs pressure for 15 minutes.

2.      Then, the flask was aseptically inoculated with 5 ml of overnight E. coli culture and kept in a shaker for constant aeration and agitation.

3.      The flasks were removed at an interval of I hour and samples transferred aseptically to a clean cuvette using sterile pipette.

4.      Optical density was read using a spectrophotometer at 600 nm and readings were tabulated.

5.      The readings were then plotted on on a graph sheet with Optical density values representing growth on the Y-axis and incubation time on the X-axis.

Observation and Result

            The optical density values for each hour time interval was tabulated.  Growth curve was plotted with OD at 600 nm on Y-axis versus incubation time on X-axis. Generation time of E. coli was calculated by extrapolation from the plotted curve.

GT = t _OD0.4- t _OD0.2

GT = 180 minutes – 150 minutes = 30 minutes

Observation (on left-hand side)

Time (Hours)	Optical Density at 600 nm
0	...
0.5	...
1	...
2	...
3	...
4	...
5	...
6	...
7	...
8	...
24	...