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Forests floors would be piled high with organic matter without the help of soil micro and macro organisms. Plants would not grow due to lack of nutrition, herbivores would cease to exist due to lack of food, and the beautiful outdoors we admire would not be alive and thriving. Microorganisms play a major role in the recycling of soil nutrients that allow life to exist, turning organic matter to proteins and nutrients in a form that can later be used by macro organisms. Nitrogen fixation is one of the most important processes done by certain soil bacteria as it allows for nitrogen to be recycled on this earth therefore creating a nitrogen homeostasis in the biosphere (Aquilanti et al., 2004). Soils worldwide offer a variety of microhabitats that certain microbes are adapted to with multiple factors that influence the presence of these microbes in the soil; temperature, pH levels, moisture, oxygen content, and nutrient levels are all very important (Prescott et al., 2008). This experiment was preformed in order to isolate and identify a bacterial genus from a soil sample by performing a series of biochemical and environmental tests.
Materials and Methods
This experiment was preformed over a five week period and started with the isolation of bacteria from two soil samples, forest floor collected from Keith Egger and coarse woody debris, which came from Hugues Massicotte (Robertson and Egger, 2008). These two samples were diluted to 10-2 in succession to 10-7 with deionized water. After isolation several techniques of culturing were used including slants, broths, deeps, and TSA plates (spread, pour and streak), all prepared using proper aseptic techniques. Week one each soil sample was diluted, all techniques following procedures outlined in the lab manual (Robertson and Egger, 2008). Lab one involved the isolation and culturing of bacteria using the prepared dilutions and the described culturing techniques. Lab two involved sub-culturing of four bacteria in order to obtain pure cultures. Four new TSA plates and slants were prepared and gram staining was done on each of the 4 selected bacteria. The four select bacteria were also viewed to determine bacterial colony morphology and with 1000x magnification to determine bacterial cell morphology and gram stain. Week three entailed all of the biochemical testing and determination of nutrient cycling for each bacterial colony. This was done by preparing 2 starch plates for starch hydrolysis, 4 SIM deeps for H2S reduction and Motility testing, four 4% peptone broth tubes for ammonification, 4 ammonium sulfate broth tubes and 4 nitrite broth tubes for nitrification testing, 4 nitrate broth tubes for denitrification testing and catalase testing using hydrogen peroxide and microscope plates. All these prepared tubes and plates were tested the following week, as well in week 3, 4 new TSA streak plates were prepared for each bacteria. During week 4 environmental factors such as pH, temperature, and osmotic pressure experiments were set up for each bacterial culture. Four new TSA plates were created each tested at the varying temperatures 4, 10-15, 22 and 37Û«C, as well four more TSA plates were created each with varying salt concentrations ranging from 0, 0.5, 2, 5 % NaCl. The pH was tested using 16 TSB tubes using the pH levels of 3, 5, 7, and 9 for each bacterium. Results from each test were observed during week 5.
Bacteria culture 1, grown on a TSA plate from the 10-7 dilution of the forest soil was studied and all characteristics of this bacterium are shown in table 1.
Table 1. All biochemical and environmental tests and yielded results from the unknown bacteria sample.
Principle Visual Properties
The colony had circular form, flat elevation, entire margin, dull appearance, opaque optical purity, no pigment - touch of yellow colour, smooth texture and was about 3mm in diameter.
At 1000x magnification the cells were rods (bacillus) and were clustered or single and had the dimension of 1.9 µm x 0.46 µm
Denitrification (NO3- to NO2-)
Denitrification (NO3- to NH3 or N2)
Nitrification (NH3/ NH4+ to NO2-)
Nitrification (NH3/ NH4+ to NO3-)
22 Û«C Mesophile
Optimal salt concentration
The colony morphology was observed from the agar plates of first subculture growth. The enumeration observed was 3.8x108 CFUs/g soil. A gram stain was conducted which allowed for cell morphology to be determined. The only result collected in lab 3 was a catalase test which showed positive once hydrogen peroxide was added and bubbles were formed with the release of oxygen. The biochemical test results were collected mainly in lab 4. Bacterium one was tested for ammonification by the addition Nessler's reagent to the peptone broth culture and a yellow colour appeared. The test for H2S reduction and motility showed no black precipitate in the SIM deep and there was no sign of bacterial growth away from the needle stab line. The starch hydrolysis test stained the agar plate a blue/black as the iodine was added. A nitrification test for both the conversion of NH3/ NH4+ to NO2- and NH3/ NH4+ to NO3-, was determined by the ammonium sulfate broth culture turning a dark purple colour with the addition of Trommdorf's reagent and H2SO4 and the nitrite broth cultures turning dark brown/ purple with the addition of Trommdorf's reagent and H2SO4 and changing to a blue colour with the addition of diphenylamine reagent and H2SO4. Denitrification was determined for both NO3- to NO2- and NO3- to NH3 or N2 reactions as there was no change when reagents A and B were added to the nitrate broths and the broth turned dark pink/ red when reagent C was added. During lab 5 the results were collected for the environmental factor tests. Bacterium one grew predominantly at an optimal temperature of 22 Û«C with the next best growth environment at 50 Û«C, at a pH level of 5, and an optimal osmotic pressure of 0% NaCl, though growth did occur in all salt concentrations.
Through a series of biochemical an environmental tests table 1 was established and based on those results collected from experiments 2 through 4 the identity of bacterium one was determined to be genus Azotobacter, family Azotobacteraceae, and order Pseudomonadales (Garrity, 2005).
Isolated bacterium one culture was first identified to be gram negative due to the lack of peptidoglycan in the cell walls and when viewed, cell morphology showed rod shapes with average dimensions of 1.9 µm x 0.46 µm. Our culture was catalase positive suggesting an aerobic organism. The three major results of gram stain, catalase testing, and cell shape were the first keys to identification of the unknown and all results are with in the characteristics of Azotobacter (Garrity, 2005). Azotobacter are normally found in single forms but can also been seen in irregular clumps and can be motile or nonmotile, as seen with bacterium one (Garrity, 2005). Further tests narrowed the search, with a negative test for all denitrification; this allowed for the cancellation of certain Pseudomonas, Bacillus, and Micrococcus, all of which can use nitrate during anaerobic respiration (Robertson and Egger, 2008). A key test in the identification of Azotobacter was the nitrification test as this bacteria is known for its role in nitrogen fixation in soils (Aquilanti et al., 2004), the isolated bacterium tested positive for nitrification. Due to the presence of Azotobacter in soils for nitrogen fixation it was expected that their optimal growth temperature would be around 22 Û«C, which categorizes them as a mesophile, and optimal growth is at a lower pH of 5, common for the Azotobacter in the presence of fixed nitrogen, (Garrity, 2005); this categorizes them in the acidophiles (Robertson and Egger, 2008).
Ammonification, starch hydrolysis, H2S reduction, and salt concentration results provided little evidence to the identification and could be used for further analysis in order for more accurate identification.
The process of nitrogen fixation can be largely affected by the presence or lack of carbon sources (Garrity, 2005), this could be used to prepare further tests for a more accurate identification of the bacteria or to go further and identify the species. Due to experimental error which could have occurred throughout the experiment, during aseptic techniques, heat fixation prior to staining, errors while performing biochemical tests, or minimal bacteria growth being present and not seen therefore concluding an experiment negative when it should have been positive or vice versa. As well further tests could be conducted with various soil samples as Azotobacter has a stronger presence in the irrigated soils with higher moisture content, therefore having a more controlled soil sample with known history would allow for better identification (Fuller and Hanks, 1982).
Isolation and identification were successful through a five week period though improvements could always be made and further experimentation could guarantee conclusions made.