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In soil, bacteria play a crucial role in soil formation, moisture regulation and nutrient cycling and are the main indicator of the soil ecosystem health (Verstraete and Mertens 2004, Entry 2008). The rapid lifecycle rate of microbes allows for any impacts of anthropological caused change (such as agriculture) to quickly be detected in microbial communities before biota in the soil show possible effects (Entry 2008). Soils that differ in water volume, temperature, and limiting nutrients (e.g. carbon (C), phosphorus (P) and nitrogen (N)) are favoured by different microbial communities (Entry 2008, Demoling 2007). These variables will differ in part on the plants they support (trees or crops) therefore it would be expected to find different microbe communities in these different areas (Demoling 2007. The purpose of this study is to isolate a microbe from either a forest or agricultural soil sample and determine to which genus it belonged through testing the effects of environment conditions, nutrient use and presence of different enzymes.
Refer to BIOL 203 Microbiology Laboratory Manual (Robertson and Egger 2010) for complete methods used. Microbe colonies from two soil samples (forest and agriculture) were grown on spread plates, streak plates and pour plates. Four distinct colonies were picked and pure colonies grown on streak plates. Each isolated sample was examined microscopically to verify by size if it was a bacteria or a yeast. An isolated colony that was determined to be bacteria was picked to identify.
To identify the bacterium, distinguishing characteristics were investigated. The bacterial colony and cell structure were visually examined. The size of the bacterium was measured using a compound microscope (with appropriate calibration calculations). The composition of the cell wall was determined through Gram staining. Colonies grown on a starch medium were bathed in iodine to test for Î±-amylase. In deeps, the reduction of sulfur was examined with iron salt to detect H2S. In the same deep, motility of the bacterium was observed. The presence of tryptophanase was examined by testing for indole with Kovac's reagent. Growth in peptone broth was tested for ammonification with Nessler's Reagent. The ability to oxidize ammonium (NH4+) to nitrite (NO2-) was tested for with Trommdorf's reagent and sulfuric acid (H2SO4) in ammonium sulfate broth. Growth in nitrite broth was tested for with the same reagents to examine if the bacteria oxidized NO2- to nitrate (NO3-). Growth in nitrate broth was tested with H2SO4 and diphenylamine reagent for NO2-. The bacterium's ability to reduce NO3- was tested by growing the bacterium in a NO3- broth which was then tested for NO2 with N,N-dimethyl-1-1-naphthylamine and sulfanilic acid. If none was detected, zinc powder was added to detect possible NO3-.
To examine optimal environmental conditions the bacterium was grown in varying pH (3 pH, 5 pH, 7 pH and 9 pH), salinity (0%, 0.5%, 2% and 5%) and temperature (4°C, 22°C, 37°C and 50°C). The amount of growth was visually determined for temperature and salinity discretely. Use of a spectrophotometer determined the amount of growth in the pH broths. To determine the mode of respiration, the bacterium was grown in an anaerobic environment (thioglycollate medium). Catalase was tested for by adding H2O2.
The bacterial colony that was picked was circular, umbulate with a rough surface. It was observed to be opaque with no pigments and was approximately 4 mm in diameter. The cells were observed to be approximately 3-5 Âµm single rods and Gram positive. See Table 1 for results summary for tests conducted to determine presences of characteristic enzymes, the use of nitrogen and which environmental conditions the bacterium isolate preferred.
Table 1. Summary of results of tests conducted to determine enzyme presence, nitrogen use and optimal environmental growing conditions for the unknown forest bacterium isolate.
NH4+ to NO2-
NO2- to NO3-
NO3- to No2-
NO3- to NH4+ or N2
When iodine was added to the bacterium colony on the starch medium, a white boarder formed around the colony indicating the bacterium contained Î±-amylase and capable of hydrolyzing starch. Black ferric sulfide was not formed when iron salt was added to the colony signifying the bacterium's inability to reduce sulfur. The bacterium migrated from the initial deep stab, indicating the presents of motility apparatus. No colour change was observed during the test for tryptophanase indicating indole was not produced. When Nessler's Reagent was added to the peptone broth it reacted with NH4+ producing a yellow colour confirming the bacterium oxidized the organic nitrogen. The bacterium was found to not oxidize NH4+ to NO2- shown with an orange product between Nessler's reagent (indicating NH4+ still present) and negative test results (no colour change) for NO2-. A blue colour was produced in the nitrite broth when diphenylamine reagent was added representing the bacterium's ability to oxidize NO2- to NO3-. It was also found the bacterium was capable of denitrification as a red colour was produced indicating a reaction between NO2- and sulfanilic acid and N,N-dimethyl-1-1-naphthylamine after the bacterium was grown in a nitrate broth. No further testing of the nirate broth was conducted because if further denitrification did take place, NO2- would not have been present. The greatest amount of colony growth was observed at 37°C, 5 pH and at 0% and 2% salinity. Oxygen bubbles were observed when H2O2 was added indicating the catalase presences. When grown in thioglycollate medium, growth was only observed along the top of the tube indicating it being an obligate aerobe.
The lack of pigment in the colony discounted a large number of genus' and was confidently determined. Other tests with clear results such as the hydrolysis of starch, motility and ammonification were next taken into consideration. The Gram stain test results were not heavily weighed as some genus contain both Gram positive and negative bacterium and there was a higher degree of human error with incorrect alcohol washing duration. The summary table of commonly found soil bacterium prepared by Eggar (2010) was used. Once the experimental tests were taken into consideration (discounting the results of the nitrification/ denitrification tests) four possible genus' were determined: Azospirillum, Pseudomonas, Acetobacter, and Bacillus. The unknown bacterium was found to denitrify NO3- to NO2- but to not denitrify NO2-. Members of Acetobacter do not denitrify NO3- while Azospirillum and Pseudomonas do denitrify NO2-. Therefore the unknown bacterium is thought to be from the genus Bacillus.
This finding is not surprising given the commonality of Bacillus ssp. in soil (Travers, Martin and Reichelderfer 1986 and Priest). It is part of the family of bacteria that form spores (Nicholson 2002). In the soil, the isolate breaks down nitrogenous organic compounds into ammonium which is then available for other biota to oxidize. This bacterium is a heterotroph as it hydrolyzes starch for a carbon source. As an obligate aerobe it uses O2 as the final electron acceptor in it's electron transport chain and produces CO2.
Another technique could have been used to confirm bascillus as described by Travers, Martin and Reichelderfer (1986) in "Selective Process for Efficient Isolation of Soil Bacillus Spp.". They selectively inhibited Bacillus ssp. by washing the soil sample with sodium acetate then allowed the sporeformes to germinate. The sample was then heat treated to kill all nonspores and germinated sporeformes leaving a pure sample of spores that were subsequently grown.
Sources of error in this experiment may have arisen during the Gram staining. The amount of time the alcohol bath was performed would change the results of the test. If the alcohol was not left for long enough Gram negative species would have still retained the outer membrane of the cell wall, allowing the thin layer of peptogylcan to be unaltered and retain the crystal violet thus staining positive.
The objective of this lab was reached. The identity of the unknown bacterium colony was determined to Bacillus via: cell morphology; cell size, shape and wall type; presence of flagellum; presence of enzymes Î±-amylase, catalase and extracellular proteolytic enzymes allowing the bacterium to break down starch, H2O2 and proteins respectively; denitification of NO3- and nitrification of NO2-; respiration mode; and optimal environmental conditions. Collectively these results rule out all other common soil microbes considered, concluding the probable microbe to be from the genus Bacillus.