Microbial communities in soil are an essential component of terrestrial ecosystems. They are crucial to the process of nutrient cycling as they decompose dead organic matter, releasing nutrients that can be incorporated into plant tissue, ultimately regulating the productivity of an ecosystem through mineralization (Buckley and Schmidt 2003). In nutrient poor environments, nitrogen-fixing bacteria and micorrhizal fungi are important to plant survival as they are involved in the acquisition and supplementation of limiting nutrients (i.e. N and P) that are otherwise unavailable to plants. Soil microbes are also fundamental to determining plant species richness by controlling plant abundance through microbial pathogens and symbiotic relationships (van der Heijden et al. 2007).
Soil provides habitat for an extremely diverse array of microbial species including procaryotic archea and bacteria, protozoa, nematodes, eucaryotic and micorrhizal fungi, and algae (Young and Crawford 2004). Soil microbes are distributed relative to the microenvironment of the soil matrix, inhabiting patches depending on clay content, moisture profile, substrate availability, acidity, and temperature (Young and Crawford 2004). Environmental factors are variable, leading to the immense diversity of microbes and conditions in which they can successfully survive and function.
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Forest and agricultural soils can differ in their microbial community profile and the conversion of natural soils to agriculturally managed soils can lead to shifts in bacterial community structure (Ovreas et al. 1998). It is therefore of essence to study the microbes, namely bacteria, potentially found in soil to determine the effects of human agricultural practices on microbial activity in soil. The purpose of this study is to isolate and identify an unknown soil bacterium from a forest soil sample to genus and master the techniques and biochemical tests involved in soil bacteria identification. The techniques and analysis tools of this study can then be applied to further bacterial identification in researching differences in forest and agricultural soil bacteria profiles.
Bacterial colonies were aseptically extracted from agricultural and forest soil solutions. Culturing and sub-culturing of the bacterial isolates were conducted in a sterile manner. A forest soil bacterial colony was chosen and examined macroscopically and observations for key, distinguishing traits were made. Individual bacteria from the colony were analyzed microscopically under oil immersion (1000X magnification) to determine the bacterium`s cellular characteristics.
The forest bacteria were subjected to the Gram staining procedure to determine their cell wall composition (Gram positive or negative). The colony was examined for motility within its growth medium. A series of biochemical tests were then conducted to determine the metabolic and enzymatic functions of the bacteria which included tests for starch hydrolysis, H2S reduction, indole production, ammonification, and two different variations of both nitrification and denitrification. Growth of the forest soil bacteria in thioglycollate medium was examined to determine their oxygen requirements. The colony was tested for the presence of catalase and an oxidation enzyme after all the other biochemical tests were completed. Growth at varying pH, temperature, and salt concentration was examined to determine the bacterium`s optimal environmental growth conditions. Refer to the laboratory manual for further detail concerning biochemical testing, environmental factor analysis procedures, and aseptic techniques (Robertson and Egger 2010).
Table 1 provides the macroscopic observations of colony morphology and the microscopic observations of cell morphology. The gram stain revealed that the bacterium retained the Safranin counter-stain. In the starch hydrolysis, the agar turned a blue/black colour. No black precipitate was noted in the H2S reduction and when observed by the naked eye, the colony showed no spread from the initial stab line. The addition of Nessler's reagent for ammonification caused a pale yellow colour change. The indole production test resulted in a yellow colour change. The denitrification of nitrate to ammonium/atmospheric nitrogen produced a red colour change. No reaction was observed for the conversion of nitrate to nitrite. Nitrification showed no colour change for the oxidation of ammonium to nitrite, though the oxidation of nitrite to nitrate showed a dark blue colour change. Bubbles formed after the addition of H2O2. The growth of the bacterium was noted throughout the tube, though was more substantial near the top. The forest bacterium demonstrated an optimal temperature of 22°C. The bacterium was most successful at pH of 7 and a salt concentration of 2%. See Table 1 for biochemical test results and classification with respect to environmental conditions.
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Table 1. Test results for unknown forest soil bacterium
Circular, slightly convex, entire, shiny, translucent, yellow, smooth, 3 mm in diameter
Bacillus (rod), singular and random arrangement, ~1.0Î¼m x 2.0 Î¼m
(NO3- to NO2-)
(NO3- to NH4+ or N2)
NH4+ to NO2-
NO2- to NO3-
Optimal salt concentration
The results in Table 1 provide strong evidence for the identification of the unknown forest bacterium as genus Flavobacterium. According to Ratner (1984), Flavobacterium have been verified as Gram negative, rod-shaped bacteria that form colonies with a distinct yellow pigmentation, and that appear smooth, translucent, entire and shiny. The bacteria were confirmed Gram negative by the staining procedure and observations of cell and colony morphology recorded in Table 1 correspond to a substantial degree with literary descriptions of Flavobacterium.
A soil dwelling Flavobacterium extracted and identified by Yoon et al. (2006) was found to grow optimally at a pH of 7 and a temperature of 25°C, optimums that are parallel with the values documented in Table 1. In addition, the bacterial colony in this study was measured to be 3 mm in diameter after 48 hours of incubation. Flavobacterium colonies were measured to be 1-1.5 mm in diameter after 24 hours of incubation (Ratner 1984). The colony size of the unknown thus may have doubled with the extended time of incubation in this study. The bacterial colony demonstrated growth throughout the tube though more near the top in the oxygen profile test, prompting its classification as a facultative anaerobe. According to Holmes et al. (1984), however, Flavobacterium are strictly aerobic although may be mistaken as facultative anaerobes as they are able to grow anaerobically in the presence of 7% carbon dioxide.
The bacterial isolate was discovered to lack the ability to hydrolyze starch, demonstrated no motility, and did not produce H2S or indole. The bacterium did not denitrify nitrate to nitrite nor did it nitrify ammonium to nitrite. The isolate was however confirmed to be capable of ammonification, denitrification of nitrate to ammonium/atmospheric nitrogen, and nitrification of nitrate to nitrite. Catalase was found to be produced by the bacterium. Flavobacterium demonstrated similar biochemical test results, though varied in nitrifying and denitrifying abilities (Holmes et al. 1984). The nitrification procedure was determined to be of low accuracy due to the variability of bacterial strains. Mutations that confer the ability to nitrify may have been acquired over time in different strains of soil bacteria. Therefore, it is still likely that the unknown belongs to this genus.
Other tests that could strengthen evidence for the choice of genus would be to test for the presence of Deoxyribonuclease and phosphatase production (Flavobacterium test positive), as well as KCN tolerance and Malonate utilization, both for which Flavobacterium test negative (Holmes et al. 1984).
The biochemical tests performed in this study were limited in terms of the observational perspective as some colour indicators produced rather lightly coloured products, such as ammonification, that could be interpreted differently. Colony and cell morphology observations were slightly objective and individuals with more experience in bacterial identification may make more educated observations. This study was also limited in that it occurred in a laboratory environment with constant environmental variables; those tested would fluctuate in a natural soil environment. The study was also confined to a certain number of biochemical tests and the accuracy they provided.
Possible sources of error are potential contamination of the colony examined by other strains of bacteria growing nearby on the agar medium, lack of sterility during culturing and sub-culturing procedures (not enough proximity to the flame), and dripping of condensation onto bacterial cultures due to improper handling of agar plates.
Some species of Flavobacterium have been linked to degrading chlorinated phenols (CP) in soil, preventing build-up of CP compounds to toxic levels (Steiert et al. 1987). Such strains of Flavobacterium thus may occupy natural soils that have been polluted by industry or that contain chlorinated phenols due to natural processes. This aspect of the bacterium may have implications for further research in bioremediation. Some members of genus Flavobacterium can perform denitrification and ammonification (Vymazal et al.1998) and thus the bacterium may play a role in the global nitrogen cycle and the release of nitrogen compounds into the soil matrix for plant utilization.
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The biochemical tests and observations made for the unknown forest soil bacterium provide strong evidence, though not complete certainty, that the unknown belongs to the genus Flavobacterium. The unknown bacteria was thus identified to the genus level, and could be further determined at the specific level with genetic and molecular analyses. The techniques and biochemical tests performed in this study could be applied in identifying numerous other soil microorganisms to determine the microbial communities of forest soils and comparing them to agricultural soils.
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