ABSTRACT

The present study was done in October 2008 and March 2009 to explore the seasonal effects on soil physico-chemical properties and fungal population found at different depth (that is, 5, 10 and 15 cm) as well as on plant parts. Results indicate that in both seasons, organic carbon, nitrogen, phosphorus and potassium was higher in the top 5 cm followed by lower depths (10 and 15 cm). Fungal species isolated from different soil depths indicate a decline in number of fungal species with an increase in soil depth, except in the rhizospheric region where the number of fungal species was higher due to highest microbial activity in this region and higher nutrient contents (due to degradation of organic compound into simpler one). In case of isolation from plant parts, it indicates that the population of fungal sp. was higher in root region compared to shoot region.

Key words: fungal species, physic-chemical properties, organic soil.

Collection of soil samples

 

Soil and plant samples were collected from the site in October 2008 and March 2009 seasons. Samples were used for physico-chemical characteristics of soil as well as the isolation of fungal organisms from both soils as well as plant parts. Samples were collected in sterile zipper polyethylene bags and stored at 4°C before processing.

 

Physicochemical characterization of soil samples

 

Physicochemical parameters include organic carbon/nitrogen, pH, water content and temperature etc. Microbial population density generally decreases with depth as a function of the availability of organic carbon and molecular oxygen, parameters which typically decrease with depth. Temperature of the soil samples was recorded on the spot. Soil samples were dried at 60°C for 72 h, powdered in pestle and mortar and filtered through 2 mm sieve and the sieved soil moisture content, pH, % organic carbon and % organic nitrogen were measured according to standard procedure as next explained.

 

Soil moisture content

 

Moisture content was determined by weighing the field fresh soil samples separately followed by drying the soil in an electric oven and then reweighing. The loss in weight is expressed as percentage of oven  dried weight.   The  samples  were  dried in  an oven at 105°C for 24 h at constant weight then cooled in desiccators and then moisture content was calculated by the formula.

 

Soil pH

 

20 gm soil was  taken in 100-ml beaker to which 50 ml distilled water (1:2.5(2.5w/v) was added and vortexing for 5 min at 120 rpm then pH was measured by digital pH meter (Orion720A+).

 

Electrical conductivity

 

20 gm of soil was shaken immediately with 50 ml distilled water in a beaker for one hour and allowed to stand overnight. Alternatively, the clear extract, after pH determination, was used for electrical conductivity.

 

The conductivity of the supernatant liquid was determined with the help of salt (conductivity) bridge meter. A 0.01 M KCl solution was used for standardization of conductivity meter before taking the sample reading.

 

Organic carbon and organic matter

 

Soil (0.1 g) was mixed with 10 ml 1 N K₂Cr₂O₇ and 20 ml of H₂SO₄. The flask was allowed to stand for 30 min. Then 5 ml of concentration orthophosphoric acid and 25 ml distilled water was added and allowed to cool. Thereafter, two drops of diphenyl amine indicator was added. Then contents were titrated with 0.5 N ferrous ammonium sulfate. At the end point solution turned into green color representing the percentage of organic carbon present in a soil sample. A blank without soil run by the same procedure (Walkley and Black, 1934). For calculating organic matter, the van bemmelen factor of 1.724 is used because organic matter contains 58% carbon.

 

Nitrogen estimation (alkaline permagnate method)

 

In an 800-ml dry kjeldhal flask, 20 gm of soil was taken. To this, 20 ml of water was added followed by 100 ml each of the 0.32% KMnO₄ and 2.5% NaOH solution. To prevent frothing and bumping, two drops of liquid paraffin was added. The contents were distilled in a collected assembly at a steady rate and the NH₃ liberate collected in a 250-ml conical flask containing 20 ml of 2% boric acid (mixed with methyl red and bromocresol green). By the absorption of ammonia, the pink color turns green. About 125 ml of distillate collects and titrate with 0.02 N H₂SO₄ and again turns into pink at the end point. Blank was also run without soil followed the same procedure (Subbiah and Asifa, 1956).

 

Available phosphorus

 

The soil of 2.5 gm was mixed with 20 ml 2% solution of sodium bicarbonate (NaHCO₃) and a spoon of charcoal was shaken for 30 min. After shaking, samples were filtered and 5 ml of this aliquot was mixed with 1 ml composite reagent (5 ml of 3% ammonium molybedate, 12.5 ml of conc. H₂SO₄, 5 ml of ascorbic acid and 25 ml of potassium antimony tartrate). In case of blank (control), distilled water was taken in place of aliquot. After 5 min, blue color was developed. The optical density was measured by UV Spectrophotometer at 885 nm wavelength. Various dilutions at the interval of 0.1 mg P/L was prepared from standard phosphate solution (K₂HPO₄) of 10 ppm (Olsen et al., 1954).

 

 

 

 

Available potassium

 

Five gram of soil was shaken on an electrical shaker with 20 ml 2% ammonium acetate for 30 min and filtered immediately through dry filter paper. The volume of filtrate was maintained up to 20 ml by adding distilled water. Potassium concentration in the extract was determined with the help of flame photometer. After standardization, calibration was done by solution of different concentration (10, 15 and 40 ppm) of potassium (Jackson, 1958).

 

Isolation of fungi from soil by serial dilution agar plating

 

Soil samples were collected from different depths, that is, 5, 10 and 15 cm. Ten gram soil was added to sterilize distilled water to make 10 ml. Serial dilutions of 10^-1, 10^-2 and 10^-3 was prepared by pipetting 1 ml volume into 9 ml sterile distilled water for additional dilution. Finally X10^-3 serial dilution soil: water suspension was prepared. 1 ml of this suspension was taken out and inoculated into nutrient plates. The plates were incubated at 25±2°C for 7 days and after the incubation period was over the plates were identified (Figure 1).

 

The infected root, stem and fruit samples were cut into small segments, washed thoroughly under tap water followed by distilled water. The samples were surface disinfected by putting them into 2% sodium hypochloride solution for 5 min. The samples were washed two times with distilled water and were blotted on sterilize filter paper for drying of adhered water with samples.

 

The sections of root stem and fruit samples were placed on autoclaved Czapek -Dox agar plates and were incubated at 25±2?C for 7 days. After incubation period was over, the colonies appeared were identified with the help of literatures available (Murumkar and Chavan, 1985; Saxena and Singh, 1987) (Figure 2).

 

 

On the basis of above experiments it was observed that the first sampling of the pigeon pea crop field which was done in September 2008 showed that moisture content was 8.81% at 5 cm, 10.68% at 10 cm and 9.43% at 15 cm depth while sampling done in March 2009 showed that percentage moisture content at 5 cm was 5.45%, 9.86% at 10 cm and 8.72% at 15 cm (Figure 3). The R² value was found to be 0.8 and 0.9 for first and second sampling   respectively.   pH   ranges of 6 .9  to   7.1  was observed at all depths in both sampling periods (September 2008 and March 2009).  There was not such variation.

 

 

 

 

From Figure 4, electrical conductivity ranges from 0.16 to 0.21 mmho at all the depths, that is, 5, 10 and 15 cm depth in September 2008 sampling period while E.C. ranges from 0.15 to 0.16 mmho in March 2009 at all respective depths.

 

In September 2008 the percentage organic carbon was found to be 0.58% at 5 cm depth, 0.51% at 10 cm depth, and 0.36% at 15 cm depth; while in March 2009, percentage organic carbon was 0.50, 0.45 and 0.26% at 5, 10 and 15 cm depth respectively (Figure 5). The R² value was calculated as 0.955 and 0.913 for first and second sampling respectively.

 

The percentage organic matter was 1.0% at 5 cm depth, 0.88% at 10 cm depth and 0.61% at 15 cm depth in September 2008, and in March 2009 it was found to be 0.87% at 5 cm, 0.77% at 10 cm, and 0.45% at 15 cm depths (Figure 6). The R² value of both sampling seasons was found to be 0.955 and 0.912 respectively.

 

Available nitrogen in sampling period September 2008 was found to be 263.42 kg/ha at 5 cm depth, 254.02 kg/ha at 10 cm and 214.29 kg/ha at 15 cm depth respectively; while in March 2009 its amount was 252.96 kg/ha at 5 cm depth, 237.4 kg/ha at 10 cm depth and 212.20 kg/ha at 15 cm (Figure 7). The correlation coefficient R² value was 0.925 and 0.981 at first and second sampling periods, respectively.

 

Available phosphorus calculated in September 2008 at 5 cm depth was 32.68 kg/ha, 25.91 kg/ha at 10 cm and 20.17 kg/ha at 15 cm depth. In March 2009, available phosphorus level was estimated as 27.84 kg/ha at 5 cm, 22.63 kg/ha at 10 cm and 19.70 kg/ha at 15 cm depth (Figure 8). The R² value was found to be 0.997 and 0.974 at first and second sampling periods respectively.

 

Available potassium level was found to be 228.27 kg/ha at 5 cm, 217.41 kg/ha at 10 cm and 215.78 kg/ha at 15 cm depth in September 2008; while in March 2009 its level was 227.92 kg/ha at 5 cm, 214.72 kg/ha at 10 cm, 183.04 kg/ha at 15 cm depth of soil (Figure 9). The R² value was found to be 0.845 for first sampling period and 0.946 for second sampling period.

 

During isolation of fungal species from soil of pigeon pea crop field, result showed that out of 14 fungal species (Figure 10) isolated from different depth of soil of pigeon pea crop field, nine fungal species viz. Aspergillus dutrenticus, Aspergillus fumigatus, Aspergillus niger, Chaetomium globosum, Fusarium sp., Fusarium udum, Penicillium citrinum, Penicillium chrysogenum and Rhizactonia solani were found at 5 cm; six fungal species viz,. Alternaria tenius, A. dutrenticus, Aspergillus flavus, Aspergillus sydowii, F. udum and Rhizopus nigricans at 10 cm depth; three fungal species viz., A. dutrenticus, A. niger and F. udum at 15 cm depth and seven fungal species viz., A. flavus,  A. fumigatus, A. niger, Aspergillus terreus,   F.   udum ,   P.  citrinum and   R.  nigricans     in rhizospheric region. The number of fungal species decreased with an increase in depth except in rhizospheric region where the number of fungal species was found higher.

 

 

 

 

 

 

During isolation of fungal species from pigeon pea plant parts, result showed that out of seven fungal species isolated from pigeon pea plant parts, three fungal species viz., Aspergillus sulfurius, Penicillium germinicum, P. citrinum  and A. niger, F. udum, Nigrosporium spharica were isolated from root length and root section followed by two from shoot section viz., A. niger and A. flavus  and one, that is, F. udum isolated  from shoot length (Figure 11). The result indicated that the population of fungal soap was higher in root region compared to shoot region.

 

From above study, it was postulated that the upper horizon contained more organic matter, nitrogen, phosphorus and other elements which in turn favors the growth of fungi in soil. The organic matter and nitrogen favored the growth of fungi in soil. In rainy season, the number of fungal colonies was more in comparison to summer due to higher moisture content which favored the germination of spores. The increased moisture content which may be attributed to the high microbial activity of soil rich in organic matter agreed with the findings of other workers (Chacko and Lockwood, 1996; Chin, 1967; Kruger, 1969; Wyk and Baard, 1971). The microbial activity was higher in monsoon season followed by winter while lower in summer season. The organic matter and nitrogen favored the growth of soil. The upper horizon contained more organic matter, nitrogen, phosphorus and other elements which in turn favors the growth of fungi in soil (Dwivedi and Dwivedi, 1972).

 

If such a variety of pigeon pea which was sown in winter and harvested in summer developed, that will be beneficial towards crop productivity as well as reduce the chances towards disease development. The interrelationship between changes in physical, chemical and biological characteristics of soil was further illustrated in  Abbott  and  Murphy  (2003).   When   one  factor  was changed, there may be a consequential effect on other factors and the final outcome was that the habitat of soil organisms was changed in several different ways following every type of disturbance (Abbott and Abbott, 1989). Organic matter and moisture content is essential to ensure the presence of an active microbial population in the soil. Physicochemical properties of the soils affect the density and diversity of microbes in the soil (Rohilla and Salar, 2012).

 

 

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