Use of advanced technologies has resulted in a significant increase in agricultural productivity. However, indiscriminate use of chemical fertilizers adversely affects soil productivity and environmental quality. Therefore, there is a great need to find some alternative which can not only enhance crop productivity without affecting soil fertility, but also maintain environmental balance.
Bio fertilizers offer an economically attractive and ecologically sound alternative to chemical fertilizers for realizing the ultimate goal of increased productivity. A bio fertilizer can be defined as a substance which contains living microorganisms which, when applied to seeds, plant surfaces, or soil, colonize the rhizosphere or the interior of the plant and promotes growth by increasing the supply or availability of primary nutrients to the host plant.
However, in certain cases of dead biological material is also considered as a bio fertilizer (some scientists prefer the terms “soil enhancer” or “conditioner” over bio fertilizer). In this article various experiments or protocols are discussed to know basic understanding and isolation of various bio fertilizers.
Experiment # 1. Isolation of Rhizobium from Leguminous Root Nodules:
Pre-Requirements:
Biological nitrogen fixation is dependent on the establishment of a symbiotic relationship between the legume and effectual Rhizobium strain. It has been observed through many studies that when legumes are first introduced into soils, the probability of root nodule formation is greatly reduced because of low population of compatible and effective strains of Rhizobium in the soil.
To combat this problem, appropriate strains of Rhizobium can be added by inoculation, especially where legumes have not been grown before and where there are no naturalised rhizobia. This can be achieved by inoculating the legume seed coat with a sufficiently large number of viable rhizobia of the correct strain, to give rapid and effective nodulation of that legume in the field, to improve crop productivity and soil fertility.
However, in tropical soils, legume inoculation often fails as there is no adequate native rhizobia and high levels of mineral nitrogen.
Thus there is an urgency to identify conditions where inoculation is needed, such as:
(1) Low species-specific rhizobia;
(2) Symbiotically related legume is not grown in the area in the immediate past history;
(3) Wastelands;
(4) Crop rotation between legume and non-leguminous crops;
(5) Low nitrate content in soil or acidic, alkaline and saline soil.
Selection of rhizobial strains for inoculant production depends upon its ability to:
a. Form nodules and fix nitrogen on the target legume;
b. Compete in nodule formation with populations of native rhizobia present in the soil;
c. Fix nitrogen across a range of environmental conditions;
d. Grow well in artificial media, in inoculant carrier and in the soil;
c. Persist in soil, particularly for annually regenerating legumes;
d. Migrate from the initial site of inoculation;
e. Colonize the soil in the absence of a legume host;
f. Tolerate environmental stresses;
g. Fix nitrogen with a wide range of host genotypes;
Isolation of Rhizobium from Root Nodules:
Specialized bacteria, which form symbiotic associations with specific hosts and fix nitrogen by forming special structures in hosts and/or their hosts, are called symbiotic diazotrophs.
All legume symbionts forming root nodules were classified into a single genus Rhizobium; however now there are at least 6 genera: Rhizobium, Sinorhizobium,Mesorhizobium, Bradyrhizobium, Azorhizobium and Allorhizobium which form nitrogen fixing nodules on roots and stems of different legumes. Collectively these are often referred to as Rhizobium.
Rhizobia can be isolated from the nodules by cleaning and surface sterilizing the nodules. Nodule contents are streaked on the plate containing yeast extract, mannitol agar medium, with Congo red. The indicator dye is added to distinguish containments from rhizobia. Most of the rhizobial strains absorb the red dye weakly and other bacteria take it up strongly.
Requirements:
CRYEMA medium, beakers, petri-dishes, sterile water, mercuric chloride (0.1%), ethyl alcohol, sterilized tiles, scalpel, glass rods and forceps.
Precautions:
i. Improper sterilization can result in growth of other bacteria
ii. Excess mercuric chloride can prevent growth on CRYEMA
iii. Sometimes nodules are cohabited by more than one strain of rhizobia or some other bacteria thereby two or more colonies can form from a single nodule.
Experiment # 2. Azotobacter- Isolation, Identification and Mass Multiplication:
The Azotobacter inoculums from the soil are made using the paste-plate method. In this technique, about 30-50g soil sample is mixed with 0.5-1.0 g of mannitol, 0.5 g of CaCO3, 0.12 ml of 10% K2HPO4 solution and 0.12 ml of 10% MgSO4 solution.
Additional distilled water is added to achieve a soil paste, followed by incubation at 27-30°C for 3-7days. After this brownish, glistening, oily colonies of Azotobacter are picked up and placed on Jensen medium and are characterized according to colony morphology, pigments production and acid production. Subsequently, Azotobacter colonies are cultured on N-free Liquid Growth (LG) media to find out their growth ability and are multiplied accordingly using the same culture media.
Experiment # 3. Azospirillum- Isolation, Mass Multiplication and Carrier Based Inoculant Preparations:
The soil sample is scraped out from the rice rhizosphere and blotted on a dried filter paper. After that, 1 g of soil is diluted by using 9 ml of autoclaved distilled water (autoclaved at 120 °C and 1.1 kPa pressure for 15 min). Then, it is successively diluted up to 10-5 level.
From the resulted 10-5 dilution, 100-μl fraction is used for plating in Okon’s media for Azospirillum. Further, it is sub cultured for pure culture and mass multiplication to prepare specific formulations. The carrier based Azospirillum inoculants are made by using Azospirillum Nfb broth (without agar) and autoclaved charcoal powder under sterile conditions.
The farmyard manure (FYM) plus soil and FYM plus charcoal or peat in 1:3 have also been used for carrier based inoculum preparation of Azospirillum. 1 kg inoculant in 40 litres of water is used for effective seedling treatment by the root dip method.
Experiment # 4. Isolation of Blue Green Algae from Water and Soil Samples:
i. From Water Bodies:
Isolation of blue green algae can be done from natural water bodies or from artificially collected water. For the isolation of blue green algae, suspend a loopful of algal growth in 5ml of sterilized distilled water, homogenize and surface-plate, 0.5 ~ 1.0 ml of it on agar plates containing a suitable medium for growth.
Isolated colonies observed through a binocular microscope after incubation are picked up, examined for contamination and transferred to test tubes containing agar slants.
ii. From Soil Samples:
Soil samples from 8-10 spots are collected from 0.5 ha-1 area after removing upper 1 cm soil crust. The samples are dried, powdered and collected and about 100gis preserved in a polythene bag for isolation purpose after thorough mixing.
There are many compositions of media for cultivation of blue green algae under laboratory conditions such as Fogg’s medium and BG-11 medium. Growth of BGA should be observed on daily basis. Colonies on growth medium can be divided on the basis of dominant, very common, common and rare. Individual colonies are lifted using an inoculation needle and suspended in 5 ml. distilled water and homogenized.
A 0.5 ml of this sample is poured on an agar plate. The plate is then rotated giving circular movement to algal suspension and incubated in light, first in an upright position for 1 day, and then in an inverted position. This process should be repeated until pure strains of algae are obtained.
Experiment # 5. Production of Azolla in Trays:
Azolla can be maintained in trays on soil-based continuous cultures. The plastic trays of dimension 45 x 30 x 12 cm may be used. Put 1 g of garden soil and tap water in the tray so that the depth of water is 5 cm. Leave it overnight so that suspended matter settles down. Inoculate with 5 g of healthy and fresh Azolla fronds.
Add tap water on alternate days to maintain water level. Add a pinch of Single Super Phosphate (SSP) if required or if fronds are showing P deficiency symptoms. Within 2 weeks a healthy mat of Azolla will cover the surface. Trays used for cultivation should be opaque in order to prevent algal growth. Regular trimming is in order to prevent over growth.
Azolla from culture trays can be applied in the field as green manure or as a dual crop along with rice (in dual cropping Azolla is grown along with rice). Fresh Azolla is inoculated @1 ton/ha and a water depth of 2 inches is to be maintained. SSP is applied @ 25-50 kg/ha in split doses. After 2-3 weeks a thick mat of Azolla is found and then rice can be transplanted.
This accounts for 10-20 ton Azolla contributing 20-40 kg nitrogen/ha. In case of pest or insect attack Furadon @2-3 kg/ha can be applied. Inoculate fresh Azolla @0.5 to 1.0 ton/ha after 7-10 days of transplantation of rice. SSP is to be applied @20 kg/ha in split doses. The incorporated Azolla dies within 8-10 days and releases nitrogen.
Another crop of Azolla should be raised during the crop cycle of rice. Each crop of Azolla during dual cropping contributes 30kg nitrogen/ha on an average.
The optimum environmental factors for Azolla production are as follows:
i. Temperature- 25-30°C
ii. Light- Partial shade
iii. pH- 4.5-8.0
iv. Moisture-Minimum level of water should be maintained
v. Phosphorous- >25ppm
Experiment # 6. Preparation of Seaweed Liquid Extract:
Take 1 kg of finely chopped seaweed and boil with 1 litre of distilled water for 1 hour in water bath. Filter the extract through muslin cloth. Allow the filtrate to cool at room temperature.
Thereafter filter through Whatman filter paper No. 41 (pore size 20-25 im). Consider the filtrate as 100% seaweed extract.
Different concentrations of Seaweed Liquid Extract; SLE (5, 10, 20, 30, 40 and 50%) can be prepared by diluting this extract with distilled water. The seaweed extract can be stored at 4°C for further applications.
Experiment # 7. Isolation of Arbuscular Mycorrhizal Fungal (AMF) Spores from Soil:
The most widely used method for isolating spores of AMF is the Wet Sieving and Decanting method. It was originally developed by Gerdeman and Nicolson (1963).
The method is as follows:
i. Prepare soil suspension in water in 1:4 ratio (w/v). Take 50 g of soil sample and mix in 200 ml of distilled water.
ii. Wait for the heavy particles to settle down and then pass the soil suspension through descending set of sieves (400- 72 number or 400 im to 25 im size).
iii. The debris retained in sieve is re-suspended in water, stirred and again passed through the 1mm sieve.
iv. This procedure is repeated until the upper layer of soil suspension is transparent.
v. Collect the content of each sieve in a petridish with the help of a water jet.
vi. Observe the suspension in petridishes with stereo- microscopes for fungal spores. The spores can be picked by flattened needle or Pasteur pipette.
The retained material on the sieve is decanted into a beaker with a stream of water and estimation of spores is carried out by modified method of Gaur and Adholeya (1994).
The Petri dish is observed under stereo binocular microscope. For the identification of AMF fungal spore, single spore or sporocarps were easily picked up from the filter paper with the help of syringe or fine point camel brush and mounted on a glass slide with a drop of polyvinyl lactophenol (PVL) and a cover slip is placed.
Subsequently, recovered spores are identified with the help of manual and different taxonomic keys proposed by different workers.
The following characters are considered for identification- sporocarps, spore morphology, size, shape and peridium of spore, sporocarps colour, wall ornamentation, subtending hyphae and mode of attachment.
Evaluation of AMF Colonization:
Arbuscular mycorrhizal fungal structure in roots is usually not observed without appropriate staining. Freshly collected root samples should be washed gently and be free from soil particles. Ultrasonic treatment is effective to disperse soil particles closely adhered to roots.
Roots were treated with 10% KOH solution for 30 min to 1- 2 hours in a hot bath, depending on thickness of root structure. Roots were washed with water and treated with 2% HCl solution. Acidified root samples are stained with 0.05% trypan blue (or acid fuchsin) in lactic acid for 10-15 min in a hot bath or for a few hours without heating.
The roots are distained with lactic acid or lacto-glycerol and are now ready for microscopic observation. The stained roots may be observed first under a dissecting microscope with transmitted illumination and then observed under a compound microscope. Fungal structures are stained blue and can be easily recognized.
Root Colonization:
Percentage of AM colonization was estimated by microscopic examination at 10 X magnification, after clearing of roots in 10% KOH and staining with 0.05% trypan blue in lactophenol according to the method described in Phillips and Hyman (1970).
The mycorrhizal colonization was determined by using the following formula:
Experiment # 8. Vermicomposting of Different Types of Waste Using Eisenia Foetida:
Vermicompost can be produced in any place with shade, high humidity and moderate temperature. Abandoned cattle shed; poultry sheds or unused buildings can be used. If it is to be produced in an open area, a shady place is selected.
A thatched roof or fiberglass cover should be provided to protect the vermicompost unit from direct sunlight and rain. The waste heaped for vermicompost production should be covered with moist paddy straw.
The various steps of vermicomposting are explained below:
Steps of Vermicomposting:
1. Collection of biodegradable wastes (crop residues, weed biomass, vegetable waste, leaf litter, hotel refuse, waste from agro-industries and floral waste).
2. Pre digestion of organic wastes for 20 days is by heaping the material along with cattle dung slurry. This process partially digests the material and makes it fit for earthworm consumption. Wet dung should be avoided as it releases a lot of gas.
3. Preparation of vermiculture bed. A concrete base is required to put the waste for vermicompost preparation. Place bedding material such as paddy straw. Put a 4 to 5 inch thick layer of cattle dung on bedding material followed by 3 to 4 inch thick layer of biodegradable wastes. Over this material, the selected earthworm (Eisenia foetida) is placed uniformly. For one- meter length, one-meter breadth and 0.5-meter height, 1 kg of worms (1000 Nos.) is required.
4. 60% moisture should be maintained throughout the period by sprinkling water occasionally. Watering should be stopped before the harvest of vermicompost.
5. Collection of earthworm is done by sieving the composted material. The fully composted material is collected and stored in bags. The partially composted material is again put into the vermicompost bed.
6. Vermicompost should be stored in shady area to maintain moisture and to allow the beneficial microorganisms to grow.
i. The depth of vermicompost pit should not be more than 3 feet because the earthworms’ activity is confined to 2 feet depth only.
ii. Do not cover vermicompost beds/heaps with plastic sheets because it may trap heat and gases.
iii. Do not overload the vermicompost heap to avoid high temperature.
iv. Maintain around 40-60% moisture. Dry conditions kill the worms and waterlogging drives them away.
v. Addition of higher quantities of acid rich substances such as tomatoes and citrus wastes should be avoided.
vi. Organic materials used for composting should be free from non-degradable materials such as stones, glass pieces, plastics etc.
vii. Use vermicompost within a month. As the vermicompost becomes aged, there is a reduction in the moisture level leading to a reduction in microbial population and activity. Consequently reduced microbial activity leads to reduced enzyme activity and NPK contents.
Chemical Analysis in Laboratory:
Use vermicompost samples at different intervals i.e. 0, 20, 40, 60, 80 and 90 days. The 0 day refers to the time of initial mixing of waste with cattle dung and soil before preliminary decomposition. Remove earthworms manually before the experiment.
Solution of vermicompost can be made by mixing lg of compost in distilled water in the ratio 1:5. Filter the mixture through Whatman paper and use filtrate for pH and nutrient analysis (Table 8.1). Determination of pH can be done by a digital pH meter, or pH strips. Following table can be used for nutrient test.
Experiment # 9. Test for Quality of Compost:
Testing compost is important because many consumers need to know the characteristics of a product before using it. Testing is also important to producers who want to use results to monitor changes in the compost product.
Testing can also help the producers, to address customer questions or complaints. The various tests carried out to study the quality of compost are given in Table 8.2.
Bio-Control:
There are several guidelines for organic farmers for using the different bio control agents.
For instance:
(a) Pheromone Traps should be installed at the rate of 10-20 traps per hectare. The distance between the traps fitted with lures is specific for a particular moth.
These traps should be positioned 6to 9 inches above the crop canopy level. Pheromones are replaced in the traps after every three weeks.
(b) Liquid formulations of NPV, Pseudomonas florescence, Bacillus subtilis and Trichoderma viridi should be kept away from direct sunlight, in moderate temperature and in well ventilated rooms. These bio pesticides should be used within the span of their specified shelf life.
It should be the endeavor of scientists to educate and create awareness about bio control agents to farmers so that they are not cheated by fake products that are sometimes sold in the market.