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The soil ecosystem includes species with unique life histories that make up a complex food web. The ecosystem's energy pyramid is founded on organic residues and consists of primary, secondary and tertiary consumers. This report focuses on bacteria found within compost soil in Prince George, British Columbia. The bacteria belong to the primary consumers which also includes a variety of fungi and simple invertebrates. The primary consumers support the ecosystem from the bottom up by feeding on detritus and organic residue and providing nutrition for organisms at higher trophic levels (Trautmann).
Bacteria, along with fungi, are paramount in the decomposition and recycling of key nutrients found in macromolecules. Most fungi bacteria are chemoheterotrophs and, therefore, use existing organic material as a carbon and energy source (Campbell and Reece, 2002, page 533). As such, they are incredibly important in the decomposition of detritus. Soil microbes also play an integral role in sulfur recycling. Some are capable of catabolizing sulfur-containing amino acids or reducing inorganic sulfur-containing compounds into hydrogen sulfide (Wind & Conrad, 1995). Many microbes, including mycorrhizal fungi and nitrogen-fixing bacteria, fix nitrogen in order to make it available for uptake by plants (Jasper, 2007). They are capable of biologically converting inert atmospheric nitrogen (N2) into nitrogen compounds (such as ammonia, nitrite and nitrate) which are then used by other organisms (Scott et al., 2008). Ammonium ion (NH4+) and nitrate ions (NO3) are then available in the soil for the plants to absorb through their root hairs. Additionally when a plant dies or an animal excretes, bacteria, or in some cases, fungi, convert the organic nitrogen in the detritus back into ammonia; this process is called ammonification (Hayatsu et al., 2008). In short, bacteria are integral to the recycling of many nutrients.
The goal of this experiment was to first isolate a bacterial culture from a sample of nutrient-rich compost soil and then identify the isolate. Several characteristics were tested in order to properly identify a bacterial isolate. Microbiologists often first gram-stain an unknown isolate to determine whether it is gram positive, having simpler walls with large amount of peptidoglycan, or gram negative, walls structurally more complex with less peptidoglycan (Campbell and Reece, 2002, page $$$). Further identification is based on an isolates particular morphology, metabolism and optimal growing environment.
Water was added to one gram of compost soil and a 10-7 dilution was prepared. One milliliter of the dilution was placed onto the bottom of an empty Petri plate and was filled and swirled with melted TSA (Trypticase soy agar) to make a pour plate. The plate was then cultured in an incubator for a week (~48hours at 25Â°C and ~120 hours at 4Â°C). A bacteria colony was chosen arbitrarily from the Petri dish and described quantitatively and qualitatively.
In order to identify the bacterial isolate, several tests were conducted to better understand its morphology, metabolism and optimal growing environment. First, a streak plate was prepared by sub-culturing a portion of the colony; this further propagated and purified the colony. Another portion was gram-stained and viewed under the microscope. The following week, the metabolism of the bacterial isolate was then evaluated. The bacterial isolate was run through the following tests: Starch Hydrolysis, H2S and Motility, Ammonification, Nitrification, Denitrification and Catalase. Each test followed the procedures outlined in the laboratory manual (Egger, 2008).
In order to determine the optimal growing environment, the bacterial culture was cultured at various temperatures, pH levels and osmotic pressures. Bacterial isolates were streaked and cultured for 36 hours on 4 TSA plates at various temperatures: 4Â°C, 10-15Â°C, 22Â°C and 50Â°C. Bacterial isolates were also cultured for 36 hours in 4 TSB tubes at various pH levels: 3, 5, 7 and 9. Finally, bacterial isolates were streaked and cultured for 36 hours on TSA plates at different salt concentrations: 0%, 0.5%, 2% and 5% NaCl.
The isolated colony from the pour plate of the 10-7 dilution proved to be circular and pulmonate in form and elevation with entire margins. It is dull, opaque and bright yellow in colour. The colony measures 1.5mm in diameter and is smooth in texture. The individual cells are rod-shaped, measure ~1.2Î¼m in diameter and are arranged in irregular chains (streptobacillus). The cells dye pink during gram-staining suggesting that they are gram negative.
Table 1.0: Starch Hydrolysis, H2S production, Motility and Ammonification Tests
H2S and Motility
Dark red/ purple
No colour change
Cloudy white media around bacteria
Pale yellow peptone broth with Nessler's reagent
Positive or negative result
Negative H2S production
As noted in table 1.0, the bacterial culture turns dark red to purple during the starch hydrolysis test suggesting that the bacteria are capable of hydrolyzing starch. This implies that the bacteria is chemoheterotrophic and uses existing organic compounds as a source of carbon and energy. The bacterial culture turns cloudy white and growth deviated from the stab line during the H2S and motility test suggesting the bacteria are motile (flagellated) but cannot degrade proteins and other sulfur containing compounds to H2S.
Table 1.1: Nitrification Tests
Ammonium sulfate broth with Nessler's reagent
Ammonium sulfate broth with Trommsdorf's reagent and H2SO4
Nitrite broth with Trommsdorf's reagent and H2SO4
Nitrite broth with diphenylamine reagent and H2SO4
"+" or "-" result
Table 1.1 reveals that the culture turns pale yellow when added to Nessler's reagent during the ammonification test indicating the bacteria are capable of catabolizing a small amount of proteins to amino acids, which are enzymatically deaminated which, in turn, releases ammonia. When the ammonium sulfate broth is inoculated with the bacteria, it expresses positive results with the Nessler's and Trommsdorf's reagent. This suggests the bacteria are capable of oxidizing ammonia to nitrite however the process is not immediate and not all the ammonia is converted. When the nitrite broth is inoculated with the bacteria, it expresses slight positive results. This suggests either only some nitrite is oxidized to nitrate or all nitrite is oxidized to nitrate but some nitrate is reduced back to nitrite. Both scenarios would explain why nitrite and nitrate are present.
Table 1.2: Denitrification and Catalase Test
Blood red broth after addition of sulfanilic acid and N, N-dimehtyl-1-1-naphthylamine
(Zinc powder was not added)
Bubbles when H2O2 was added
Positive or negative result
As noted in table 1.2, the culture turns red after the addition of sulfanilic acid and N, N-dimehtyl-1-1-naphthylamine. This suggests the bacteria are able to reduce nitrate back to nitrite using the enzyme nitrate reductase but unable to further reduce nitrite to ammonium ion. Furthermore, when hydrogen peroxide is added to the culture, oxygen is released suggesting the bacteria are catalase positive and can break down hydrogen peroxide into water and free oxygen.
Table 1.3: Optimal Temperature, pH and Osmotic Pressure
Osmotic pressure (%NaCl)
Table 1.3 shows at which temperature, pH levels and osmotic pressure the bacterial isolate grows optimally. The bacterial isolate grows best at 22Â°C at a neutral pH on 0% NaCl medium. This suggests the bacteria can be classified as mesophiles, neutrophiles and nonhalophiles.
The bacterial isolate was able to be identified after assessing several morphological and metabolic traits. The bacteria are gram negative, rod shaped, mobile and form yellow colonies. Additionally, the bacteria possess catalase which breakdowns hydrogen peroxide. These indices would suggest, according to Bergey's Manual of Systematic Bacteriology, that the bacteria belong to the family Azotobacteraceae (Krieg & Holt, 1986, p. 224). Furthermore, the bacteria fix nitrogen under normal atmospheric pressure and are less than 2Î¼m suggesting that they belong to the genus Azotobacter (Krieg & Holt, 1986, p. 226).
This particular bacterial isolate certainly fills a very important niche in the compost soil ecosystem. Compost soil is extremely nutrient rich and relies on the work of microbes to catabolize macromolecules and fix various elements. This chemoheterotrophic bacteria absorbs nutrients from dead organic matter and, in turn, decomposes the detritus and returns nutrients back to the ecosystem. Furthermore, like many other bacteria and fungi, one of the most significant roles of this Azotobacter is to oxidize ammonium into nitrite and nitrate. The latter are forms that are more easily absorbed by plants and, in turn, allows for nitrogen recycling (Campbell and Reece, 2002). Several studies have investigated the ecological role of several Azotobacter and the majority highlight the bacteria's role in nitrogen fixation (Garg et al., 2001; MunozCenteno et al., 1996; Bakulin et al., 2007). Some Azotobacter are so efficient at fixing nitrogen that they can be used as biofertilizer for enhancing nitrogen input and phosphate solubilization in fish ponds (Garg et al., 2001). Azotobacter are also known to form synergistic or additive interactions with arbuscular mycorrhizal fungi to form symbiotic relationships with plant roots (Bagyaraj & Menge, 1978). It is of no surprise to find this nitrogen-fixing chemoheterotroph in compost soil.
There are several other tests that may have aided in the identification of the bacterial isolate. Bacteria are commonly distinguished based on the presence or absence of nitrogenase, a bacterial enzyme which converts atmospheric nitrogen to ammonia (Lubambo et al., 2007). Additionally, discrepancies in growth within lighted versus dark incubation chambers would reveal any photosynthetic activity of the bacteria. Alongside the starch hydrolysis test, this would further substantiate the discrimination between heterotrophism and autotrophism. Additionally, one could evaluate the level of metabolism under various anaerobic/aerobic conditions to determine whether the bacteria are strict, tolerant or facultative aerobes or anaerobes.
There exist several sources of error and limitations to the tests that were performed. Identification was carried out by relating our observations to a dichotomous key found in Bergey's manual. The observations, however, were not strictly dichotomous but more graded. It was difficult to differentiate between a very positive and a weakly positive result when applying the observations to the key. Also, it was difficult to discriminate between a bacterium not capable of entirely oxidizing nitrite to nitrate and one that could fully oxidize nitrite to nitrate who also reduced some nitrate back to nitrite. Another contentious issue is one of purity. More than one bacterial strain may have been cultured or incorporated into the tests due to improper aseptic technique. This may have produced misleading results causing a misidentification of the bacteria.