Ampicillin and Kanamycin Resistant Bacteria Comparison

Antibiotic use throughout the world has increased tremendously over the decades. In the past, antibiotic resistance was most prevalent in areas of frequent antibiotic use, such as in medical or laboratory settings. However, the increasing use of antibiotics and antibacterial products outside of hospitals, such as in homes and schools, echoes the expansion of antibiotic resistant bacteria (LBC Biology Staff, 2010). One major source of the growing problem is that antibiotics are being over prescribed by doctors to millions of people around the world. It is currently believed that about only half of the antibiotics prescribed to patients are administered properly (Levy, 1998). In addition to over prescription by doctors, many patients misuse the antibiotics and further increase the spread of resistance. For example, some patients discontinue use of antibiotics upon feeling symptom relief, not at the end of their antibiotic schedule prescribed by the doctor. In actuality, patients are killing off the weakest bacteria, causing temporary relief, and allowing the stronger and more resistant bacteria to multiply at a faster rate (Levy, 1998). This and other types of antibiotic misuse have promoted the growth of strains of bacteria with resistance to antibiotic attack. This can be seen through studies that have shown Tetracycline resistance by normal human intestinal flora that exploded from 2% in the 1950s to 80% in the 1990s (Criswell, 2004). Other studies have shown Kanamycin, an antibiotic from the 1950s, has become clinically useless as a result of the prevalence of Kanamycin-resistant bacteria (Criswell, 2004). It has become visible that the development of resistance to any antibiotic, new or old, will happen in a matter of time (LBC Biology Staff, 2010). Due to the inevitability of mutation, natural selection, time and environmental conditions, resistance will be seen in more common areas like work and home.

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As a consequence of the every growing expansion of antiobiotic resistance, places previously thought to be uncontaminated like schools and homes have become overwhelmed with antibiotic resistant bacteria. In one household study, it was discovered that kitchen sinks contained many different types of resistant bacteria, primarily from food waste and human hands (Rusin et al., 1998). Only the application of strong bleaches and specific cleaning products on a regimented cleaning schedule led to a decreased amount of bacteria in kitchen sinks (Rusin et al., 1998). The cleaning products used in this study did not contain antibacterial ingredients, which helped reduce the spread of resistance by killing all bacteria instead of the most susceptible strains. Antibacterial products and cleaning supplies are less effective and in turn can lead to reproduction of stronger antibiotic resistant bacteria. The large amount of antibacterial cleaning products, food and waste combined with the constant water supply in sink drains allows for a greater chance of survival of antibiotic-resistant bacteria (Levy, 1998). Optimal conditions for bacterial growth with a wet environment cause a higher frequency of bacterial transmission of resistance (Perryman and Flournoy, 1980). In scientific laboratories, regulations are in place to monitor the disposal of solid and liquid wastes. Some regulations include specific waste baskets for toxic or contaminated substances and use of certain sinks only when dealing with harmful liquids in laboratory settings. This ensures that unnecessary amounts of harmful substances that could lead to resistance are not continually poured down laboratory sink drains. However, no such regulations are in effect in household environments.
In a study performed in Oklahoma City the extent of growing antibiotic resistance was seen in multiple environments. Bacterial samples were gathered from sink drains in the Veterans Administration Medical Center, libraries, private homes, shopping centers, and other similar environments for comparison (Perryman and Flournoy, 1980). The goal of the experiment was to determine the types of resistant bacteria that were most prevalent in sink drains, the abundance of bacteria in sink drains, and the life span of bacteria in dry and wet environments (Perryman and Flournoy, 1980). Through testing, bacteria were found to have longer life spans in wet environments than in dry environments, and many bacteria survived for over 180 days in wet environments (Perryman and Flournoy, 1980). The high survival rate of bacteria in areas with constant water supply, such as in laboratory and kitchen sinks, supports the prediction that sinks are ideal environments for ample bacterial growth. In the aforementioned study, bacterial growth occurred on plates containing the antibiotics gentamicin and amikacin, and it was determined that the sink drains from the medical hospital contained the highest amount of antibiotic resistant organisms. Overall, 88% of the sink drains sampled from the Veterans Administration Medical Center contained some type of antibiotic resistant bacteria (Perryman and Flournoy, 1980). While bacteria could come from other sources such as the patients and tap water, the great quantity of antibiotic resistant bacteria in all environments illustrates the need for a reduction in the overuse of antibiotics and the essential awareness of the consequences.
Places with high levels of exposure to antibiotics and antibacterial products provide ideal environments for bacteria to develop resistance through replicated mutations or transmissions between bacteria. Some factors that severely add to the growing problem of antibiotic resistant bacteria include increased applications of antibacterial soaps and cleaning products, over prescription of antibiotics by doctors, misuse of antibiotics by patients, and improper care of waste products (Levy, 1998). Bacteria can become resistant to antibiotics through genetic mutation, transfer of the mutation between bacteria, or transmission of the mutated DNA on a plasmid between bacteria when the resistant gene is carried on the plasmid DNA. A plasmid is a relatively small piece of circular DNA that is self replicating and independent of the chromosomal DNA of the cell. Resistant chromosomal DNA and plasmid DNA can be transmitted to the next generation through cell replication. Plasmids can be passed through bacterial conjugation, which involves a bacterium copying the plasmid with resistant DNA and inserting the copied plasmid into a second bacterium. Plasmid DNA can also be transferred through bacterial transformation when plasmid DNA invades another bacterium and is incorporated into the bacterium’s DNA (Cognato, 2010). Understanding these problems and the mechanisms of resistance transmission is the first step in preventing further development of resistant strains of bacteria.
The focus of the experiment at hand is to determine whether the bacteria located in a laboratory sink or in an apartment garbage disposal contains more antibiotic resistant strains. It was hypothesized that the apartment garbage disposal would contain more antibiotic resistant bacteria than the laboratory sink. This is due to the abundance of contaminated materials that pass through garbage disposals in comparison to the regulated materials that pass through laboratory sinks. The null hypothesis is that the amounts of antibiotic resistant bacteria that exist in the garbage disposal sink and laboratory sink will be equal.
Many steps were needed to accomplish this research and obtain the sample bacteria to determine the resistance. Samples from the laboratory sink and the apartment garbage disposal were swabbed on agar plates to obtain a culture of bacteria. Colonies were selected based on growth and seclusion from the bacterial “lawn”. Individual bacteria were then streaked on master patch plates for each environment. After the bacteria had grown, individual colonies were selected to be streaked on antibiotic plates containing Ampicillin, Kanamycin, and Tetracycline. Antibiotic resistant bacteria were chosen from the antibiotic plates, separated and characterized. Next, plasmids from the antibiotic resistant bacteria were isolated and spliced using restriction endonucleases to determine band length of resistant plasmid DNA to help identify the type of bacteria. Competent E. coli cells were transformed with the control plasmid DNA to convey antibiotic resistance and support bacteria identification. Finally, the bacterial DNA was replicated by polymerase chain reaction to amplify the 16S rRNA gene in hopes to obtain sequencing information of a known bacterium. It was predicted that resistant bacteria, for all antibiotics, will be Gram negative due to easier entry of resistant plasmid DNA into the cell. Bacteria with a thin cell wall layer and an outer membrane surrounding the peptidoglycan layer are Gram negative. Bacteria with a thick wall layer that do not have the peptidoglycan layer surrounding are Gram positive. Gram identity was verified through Gram staining, a KOH test, and observing growth on a MacConkey agar and Eosin Methylene Blue Agar plate.
Methods
Swab Plates
A sterile cotton swab saturated in sterile phosphate-buffered saline was used to gather samples from the laboratory sink and an apartment garbage disposal. Bacterial samples from the disposal and lab sinks were collected from the underside of the drain. Bacteria were then swabbed onto Lysogeny broth agar plates (three per environment). Plates were placed into an incubator for 24 hours at 37°C. Following the incubation period, plates were removed, parafilmed, and refrigerated at 4°C until needed.
Master Patch Plates
Master plates were made by placing sixteen individual colonies onto a 4×4 grid on Lysogeny broth (LB) only plates. An inoculation loop was used to transfer the 16 individual colonies from the sample plate onto a grid of the master plate. Plates were labeled with D for the apartment garbage disposal and L for the laboratory sink along with a number (1, 2, or 3) to distinguish between swabbed samples. Plates were incubated at 37°C for 24 hours, removed, sealed with parafilm, and refrigerated at 4°C until needed.
Antibiotic Patch Plates
Antibiotic agar plates were made by mixing 8.4g agar with 12g LB powder and 600mL of distilled water (dH2O), and then autoclaved. After cooling, 2.4µL of Ampicillin, 1.2µL of Kanamycin, or 2.4µL of Tetracycline were added appropriately and plates were poured. One colony per grid of the master patch plate was obtained with an inoculation loop, and the bacteria were transferred in a line onto a corresponding grid on the antibiotic plates. The number of squares that contained bacterial growth was observed and recorded. One colony of the bacteria grown on the antibiotic patch plates was then streaked onto a new antibiotic plate to obtain individual colonies of bacteria for further study.
Miniprep
A liquid culture was performed in preparation for the Promega Wizard Plus SV Miniprep DNA Purification System, which was used to isolate plasmid DNA from antibiotic resistant bacteria. First, 5µL of antibiotic was added to a 5mL tube filled with a liquid medium made of LB. A single colony of bacteria was added to the medium and placed in a shaker at 37°C for 24 hours. The liquid culture was then transferred into an Eppendorf tube and centrifuged for 5 minutes at 4,400rpm. Liquid media waste was disposed of and the pellet was thoroughly re-suspended in 250µL of Cell Resuspension Solution. If the bacteria were Gram positive, 63µL of lysozyme would be added to the solution. Since the bacteria studied was Gram negative, the process continued with the addition of 250µL of Cell Lysis Solution was added to the Eppendorf tube containing the resuspended bacterial solution and the sample was mixed. Subsequently, 10µL Alkaline Protease Solution was added, mixed, and incubated for 5 minutes at room temperature. Then, 350µL Neutralization Solution was added, mixed, and centrifuged for 10 minutes at 13,500rpm. A Spin Column was inserted in a Collection Tube and the clear lysate was decanted into the Spin Column. This was centrifuged for 1 minute at 13,500rpm and the flowthrough was discarded. The Spin Column was replaced, 750µL of wash solution was added, and the solution was centrifuged for 1 minute at 13,500rpm. The flowthrough was discarded, and this process was repeated with a 250µL wash. The solution was centrifuged for 2 minutes at 13,500rpm. The Spin Column was transferred to a 1.5mL Eppendorf tube. Finally, 50µL of Nuclease-Free Water was added and then the solution was centrifuged for 1 minute at 13,500rpm. The column was discarded and the DNA was stored at -20ËšC.
Gel Electrophoresis
DNA electrophoresis was used to determine the length of the plasmid DNA of the environmental samples and Blue plasmid control (pKAN). First, 0.7g of agarose powder was added to 70mL of 1X TBE. The solution was heated in a microwave for 1 minute so the agarose powder was completely dissolved. After the mixture cooled, 3µL of Ethidium bromide was added and the gel was taken out of the mold and put on the rig. The gel was submerged in a 1X TBE buffer. The wells of the gel were filled with 10µL of a mixture containing 8µL of plasmid DNA and 2µL of plasmid dye, and the gel ran for 60 minutes on 80 volts. The 1% agarose gel was viewed under an ultraviolet light to compare lengths of DNA with the 1KB ladder.
Gram Staining
Gram staining was used to determine the Gram identity of bacteria. Bacteria that are Gram negative stained red and bacteria that are Gram positive stained violet. A colony of bacteria was added to an Eppendorf tube with 400µL of dH2O. After vortexing, 5µL of the solution was pipetted onto a slide. Once dry, the slide was passed over a flame to affix the bacteria to the glass, preventing the removal of bacteria. The slide was flooded drop-wise with crystal violet and iodine, and rinsed with dH2O for 5 seconds after the addition of each reactant. Ethanol was added until the color was no longer emitted, then rinsed with dH2O for 5 seconds. Safranin was added drop-wise for 1 minute and then rinsed with dH2O for 5 seconds. The slide was observed under a microscope to determine Gram identity.
KOH Test
The KOH test for Gram positive and negative bacteria was begun by pipetting 20µL of 3% KOH on a slide. After adding one clump of bacteria to the KOH, the consistency of the solution was observed. If the solution was thick, viscous and adhered to the inoculation loop, the bacteria were Gram negative. If the solution was thin and not viscous, the bacteria were Gram positive.
MacConkey Agar Plate
A MacConkey agar plate was streaked with antibiotic resistant bacteria from the garbage disposal and laboratory sink. After incubation at 37ËšC for 24 hours, the plates were observed for growth to indicate Gram negative bacteria. The MacConkey agar plate also signaled lactose fermentation with the appearance of pink colonies.
Eosin Methylene Blue Agar Plate (EMB)
An EMB plate was streaked with antibiotic resistant bacteria from the apartment garbage disposal and the laboratory sink as well as a positive E.coli control. After incubation at 37ËšC for 24 hours, the plates were observed for growth to indicate Gram negative bacteria. The EMB agar plate indicated strong lactose fermentation through the appearance of dark green metallic colonies and a lesser degree of lactose fermentation through the appearance of purple or pink colonies.
Restriction Digest
Restriction enzymes cut the control pKAN DNA at specific restriction sites identified by the NEBcutter V2.0. The enzymes used in restriction digest were BamHI and EcoRI in Buffer II, and PvuI and PstI in Buffer III. The reaction solution used in restriction digest consists of 10µL of DNA, 1µL of each enzyme, 2µL of NEBuffer, and 7µL of de-ionized distilled water (ddH2O) added together in an Eppendorf tube. The solution was centrifuged at 14,500rpm for 30 seconds and then incubated for 24 hours at 37ËšC. A plasmid map created from the NEBcutter V2.0 was compared to a gel electrophoresis run on a 1% aragose gel with plasmid DNA. The gel electrophoresis compared Blue plasmid (pKAN) DNA that was uncut with the Blue control plasmid (pKAN) that was cut with restriction enzymes.
Transformation
After plasmid DNA preparation, 22µL of E. coli competent cells were added to three separate Eppendorf tubes. In one tube, 5µL of control DNA, pKAN, was added and stirred with the pipette tip. In the second tube, a negative control was made with the addition of 5µL of dH2O that was then stirred with a pipette tip. In the third tube, a positive control was made with the addition of 1µL of known pKAN, and the solution was stirred with a pipette tip. The tubes were then incubated in ice 30 minutes. The cells were heat shocked for 45 seconds at 42ËšC and then placed on ice for 2 minutes. 250µL of pre-warmed (37ËšC) SOC medium was added to all three of the Eppendorf tubes, and the tubes were then incubated in a shaker at 37ËšC for 1 hour at 2,250rpm. Upon removal from the incubator, 75µL of each transformation were spread onto plates with a sterilized “hockey stick”. The transformed control DNA, pKAN, cells and the negative control dH2O transformed cells were spread onto LB only plates, ampicillin antibiotic plates, and kanamycin antibiotic plates to determine if resistance to antibiotics was transferred in the transformation. The transformed positive control, known pKAN, cells was spread onto a LB only plate and a kanamycin plate since pKAN is known to be resistant to kanamycin. Plates were incubated for 24 hours at 37ËšC and numbers of resistant bacterial colonies were observed. Bacterial growth on the control DNA, pKAN, transformation antibiotic plates would signal resistance to the antibiotic in the plate, and growth on the LB only plate would signal the existence of bacterial cells from the transformation. No growth on the dH2O negative control plates containing ampicillin and kanamycin antibiotics would signal a correct transformation as long as there was bacterial growth on the LB only plate. Growth on the positive control, known pKAN, transformation plate signaled the correct transfer of kanamycin resistant plasmid DNA into the competent E.coli cells.
Polymerase Chain Reaction
The Polymerase Chain Reaction (PCR) involved mixing a reaction cocktail that included 80µL of Nuclease-free water, 10µL of 10X Thermopol buffer, 3µL of 10mM dNTPs, 2µL of 11F @ 10µM, 2µL of 1492R @ 10µM, and 1µL of Taq polymerase @ 5000U/mL. The solution was then mixed through vortexing. Subsequently, 22µL of the cocktail was transferred to each of the 4 PCR tubes. A small portion of each bacterial colony was added to SOC medium and mixed. Then 5µL of SOC medium with bacteria was added to each tube. Tube 1 had environmental bacteria, tube 2 had different environmental bacteria, tube 3 had the control E.coli and 5µL of H2O was added to tube 4. The reactions were placed in the thermocycler in C4. The PCR cycling program consists of five steps. The first step is pre-denaturation in which the PCR mixing reaction cocktail is heated at 95°C for 5 minutes. The second step is denaturation, which involves heating the reaction cocktail at 95°C for 30 seconds to unwind and separate the DNA. The third step is annealing, which is run at 50°C for 30 seconds to allow the 11F and 1492R primers to attach to the DNA template strands. The fourth step is elongation, which is run at 72°C for 45 seconds to allow the DNA polymerase (Taq polymerase) to add dNTPs and replicate the 16S gene. The fifth step is the final elongation, which is run at 72°C for 7 minutes. The hold between cycles is run at 4°C, and the PCR is run for 35 cycles. Gel electrophoresis was run to determine if a successful PCR reaction took place. 10µL of the PCR solution from each tube was mixed with 2µL of plasmid dye, and 10µL of the mixtures were loaded into the wells of the 1% agarose gel.
Chi Squared Test of Independence
A Chi Squared Test of Independence was run to determine if a statistically significant difference exists between the numbers of antibiotic resistant bacteria from the two environments. The number of grids on the antibiotic plates was recorded only if the bacteria grew on both the antibiotic plate and the LB only plate. The test was run on Vassar Stats and gave a p-value to correspond to the data and indicate if there was a significant difference.
Results
Swab and Master Patch Plates
After the incubation period of 24 hours at 37 C, the swab plates, labeled L for laboratory sink samples (L1-L3) and D for garbage disposal sink samples (D1-D3), were observed and found that 100% of the environmental bacteria grew (Figure 1). Bacteria growth in both environments was indicated by white colored spots or streaks within the plate’s grid. Master plates were observed from both experimental environments and found to have growth on all of the 16 grids on each plate (Figure 2).
Antibiotic Patch Plates
From the garbage disposal sink, the three samples all had some level of growth (Figure 3). The following percentages were calculated by dividing the number of grids with bacterial development on the antibiotic plates by the number of grids with growth from the LB plates (Table 1). Plate D1 showed 100%, 62.5%, 0%, and 100% growth on the Ampicillin, Kanamycin, Tetracycline, and LB only plates respectively. Plate D2 demonstrated 93.75%, 93.75%, 0%, and 100% growth on the Ampicillin, Kanamycin, Tetracycline, and LB only plates respectively. Plate D3 showed 93.75%, 75%, 0%, and 100% growth on the Ampicillin, Kanamycin, Tetracycline, and LB only plates respectively. From the laboratory sink, all samples had bacteria development (Figure 4). Plate L1 demonstrated 100%, 93.75%, 12.5%, and 100% growth on the Ampicillin, Kanamycin, Tetracycline, and LB only plates respectively. Plate L2 showed 100%, 73.33%, 6.67%, and 93.75% growth on the Ampicillin, Kanamycin, Tetracycline, and LB only plates respectively. Plate L3 demonstrated 57.14%, 42.86%, 7.14%, and 87.5% growth on the Ampicillin, Kanamycin, Tetracycline, and LB only plates respectively.
Chi Squared Test of Independence
Data obtained from the number of antibiotic resistant colonies on the antibiotic patch plates was used to run the Chi-squared Test of Independence for Ampicillin and Kanamycin resistant bacteria. For Ampicillin resistant bacteria, the p-value obtained was 0.74. With one degree of freedom, the Chi-squared critical value of 3.84 obtained from a Chi-squared Distribution Table in comparison to the Chi-squared statistical value denoted no statistically significant difference. For Kanamycin resistant bacteris, the calculated p-value was 0.81. With one degree of freedom, comparison of the Chi-squared critical value of 3.84 found in a Chi-squared Distribution Table and the Chi-squared statistical value demonstrated no statistically significant difference (Table 1).
Gram Staining, KOH, MacConkey Agar and Eosin Methylene Blue Agar Plates
Four tests were used to determine the gram identity of bacteria from the experimental environments. The results showed that the three environmental bacteria slides were stained pink indicating gram negative bacteria (Figure 5, Table 2). For the KOH test, all three samples from both environments appeared viscous and thick, indicating gram negative bacteria (Table 2). The MacConkey Agar Plate was divided into three sections for the different antibiotic resistant bacteria. The environmental bacterial sample in Section 1 was obtained from the Ampicillin antibiotic plate L2 grid #3. The bacterial sample in Section 2 was obtained from the Kanamycin antibiotic plate L2 grid #14. The bacterial sample in Section 3 was obtained from the Kanamycin antibiotic plate D2 grid #16. All three samples in the three sections grew bacteria that were stained pink, indicating Gram negative bacteria that ferment lactose (Figure 6, Table 2). The Eosin Methylene Blue Agar Plate was sectioned off into four parts and bacteria from three environmental samples and one E.coli positive control were plated. The bacterial sample in Section 1 was taken from the Ampicillin antibiotic plate L2 grid #3. The bacterial sample in Section 2 was obtained from the Kanamycin antibiotic plate L2 grid #14. The bacterial sample in Section 3 was gathered from the Kanamycin antibiotic plate D2 grid #16. The bacterial sample in Section 4 was obtained from an E. coli plate that was known to be Gram negative. Pink colonies formed in all four sections, signaling Gram negative identity of the bacteria and lactose fermentation (Figure 6, Table 2).
Mini Prep and Gel Electrophoresis
Promega Wizard Plus SV Miniprep DNA Purification System was run to isolate plasmid DNA. This plasmid DNA was run on a 1% agarose gel. The lengths of bands in Trial A could not be determined because the DNA in the wells did not run with the ladder. The Blue control plasmid, which was pKAN, was located in lane 3 in Trial A and Trial B and was used to indicate a successful Miniprep. The band length of the pKAN control DNA in Trial B was about 4,200 base pairs. An environmental plasmid found on Ampicillin streak plate L2, grid #3 was used in lane 7 in Trial A and lane 5 in Trial B. In Trial B, the base pair length of the environmental bacteria plasmid used in lane 5 could not be determined due to the appearance of many bands of varying length. An environmental plasmid from Kanamycin streak plate L2, grid #14 was used in lane 5 in Trial A and lane 7 in Trial B. The band length of this environmental plasmid in Trial B could not be determined due to the faint appearance of a band greater than 10,000bp. Another environmental plasmid from Kanamycin streak plate D2, grid#16 was used in lane 6 in both Trial A and Trial B. The band length of this environmental plasmid in Trial B also could not be determined from the faint appearance of a band greater than 10,000bp (Figure 7).
Restriction Digest
In Trial A, restriction digest was used to cut the Blue control pKAN DNA with the enzymes BamHI, EcoRI, PstI, and PvuI. Lane 3 displays pKAN cut with PstI and PvuI. Lane 4 displays pKAN cut with BamHI and EcoRI. The lengths of the bands shown are about 4,000bp, 3,000bp, 2,500bp, 1,500bp, and 1,200bp.The lengths of the bands shown are about 1,700bp, 1,100bp, 750bp, 600bp, and 500bp. Lanes 5-8 contained environmental bacterial DNA that was cut with BamHI, EcoRI, PstI, and PvuI as well, but no bands were observed (Figure 8).
In Trial B, restriction digest was used to cut pKAN DNA with only the enzymes BamHI and EcoRI. Lane 3 displays pKAN that was cut with BamHI, showing a band length that is about 4,200bp. Lane 4 shows pKAN that was cut with EcoRI, and the band lengths shown are about 8,000bp, 5,000bp, and 4,000bp. Lane 5 displays pKAN that was cut with BamHI and EcoRI, and the band lengths shown are about 4,100bp, 3,100bp, and 2,000bp. Lane 6 shows pKAN that remained uncut with a band length of about 4,200bp (Figure 9).
Transformation
Transformation was performed to convey resistance carried on plasmid DNA into competent E. coli cells. Blue plasmid control DNA (pKAN) was used for the transformation, which was successful. This was indicated by the growth of transformed bacteria on Kanamycin antibacterial plates (Figure 10).
Polymerase Chain Reaction
A Polymerase Chain Reaction (PCR) was used to amplify and prepare the 16S gene of rRNA. Gel electrophoresis was run on the PCR product to determine if a successful PCR reaction had taken place. Lane 3 contains PCR product from the Kanamycin plate L1 grid #14 and lane 4 contains PCR product from the master patch plate D3 grid #16. Bands were not seen in these lanes containing environmental bacteria, signaling an unsuccessful PCR. Lane 5 displays the negative water control without bands. Lane 6 shows the positive E. coli control PCR product with a band length of about 2,000bp (Figure 11).
Discussion
The study showed that no statistically significant difference existed between the amount of antibiotic resistant bacteria in the garbage disposal and laboratory sink and it also characterized all of the environmental bacteria as Gram negative. To determine the amount of bacteria located in the experimental areas, many tests were utilized to analyze the bacterium. Patch plates containing Tetracycline, Ampicillin, Kanamycin and LB were made in order to verify antibiotic resistant bacteria and growth. The plates with bacterial growth that was resistant to Ampicillin and Kanamycin were used in a statistical analysis to determine a correlation between the amounts of growth and the two environments. Our prediction that the amount of bacterial growth from the garbage disposal sink in Capitol Villa would be greater than the Lyman Briggs lab sink in C5 was refuted due to the Chi-squared Test for Independence that showed no statistically significant difference. We failed to reject the null hypothesis that no difference existed between the amounts of antibiotic resistant bacteria found in each environment.
A Chi-squared Test for Independence was run to compare the amounts of antibiotic resistant bacteria on the Ampicillin and Kanamycin plates. Tetracycline was not used because no data indicated resistance. The existence of Ampicillin and Kanamycin resistant bacteria in both the garbage disposal and the laboratory sink is unsurprising due to the widespread clinical use of both antibiotics over the past decades (Criswell, 2004). For Ampicillin, a total of 178 bacterial streaks grew between the two environments and a p-value of 0.74 was calculated. With one degree of freedom, the Chi-squared critical value of 3.84 obtained from a Chi-squared Distribution Table in comparison to the Chi-squared statistical value denoted no statistically significant difference. For Kanamycin, 162 streaks grew between the two environments and a p-value of 0.81 was calculated. With one degree of freedom, the a comparison of the Chi-squared critical value of 3.84 found from a Chi-squared Distribution Table to the Chi-squared statistical value denoted no statistically significant difference as well. Therefore, the prediction that the garbage disposal sink would contain more antibiotic resistant bacteria than the laboratory sink was rejected.
To further understand why bacteria were resistant, four tests were run to categorize the Gram identity of the environmental samples. The structure of the bacteria plays a large role in determining resistance. Importantly, it is easier for the plasmid DNA to penetrate a Gram negative bacterium due to the lack of an outer membrane around the peptidoglycan layer. The Gram staining process showed pink rod shaped bacterium, demonstrating that the bacteria was Gram negative. The KOH tests resulted in a viscous substance, indicating that all the environmental bacteria obtained from the garbage disposal and the laboratory sink were Gram negative. The MacConkey agar plates identified the bacteria to be Gram negative through growth on the plate. The growth on the plate was a pink color, signifying lactose fermentation from the bacteria. The environmental bacteria developed pink colonies on the EMB agar plates, further supporting the Gram negative identity and a low production of lactose fermentation of the environmental bacteria gathered from the garbage disposal and laboratory sink.
Gel electrophoresis was used in determining the existence and length of environmental plasmid DNA. The Miniprep isolated the plasmid DNA from the bacteria, but upon running the gel, it was discovered that no environmental plasmid DNA was present. The absence of bands
 

Plasmid Retention and Bacteria Growth in E Coli

The increasing interest by the industry in recombinant protein production has caused an intensive study in this area during the last years. However, it is well known that there are a number of issues associated with the high expression of a recombinant protein. E. coli is one of the most used organisms for this purpose. In this organism, the most common and challenging problem is the formation of inclusion bodies. Probably, an incorrect folding process provokes that the recombinant protein forms those structures. When the protein forms inclusion bodies, it is insoluble and usually useless. In order to find a proper protocol for the high production of the protein S, we have assessed the expression system which use the BL21*DE3 strain as host and the pCV05 plasmid which contains the protein S sequence fused with the His tag sequence. Growth rate, plasmid loss and recombinant expression level were assessed. We obtained a reasonable production of target protein in the insoluble fraction. Further research is needed to know whether the processing of the His tag is able to make soluble the protein from the inclusion bodies as is described by other researchers.

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Introduction
The increasing interest by the industry in recombinant protein production (RPP), due to the number of applications it can provide, has caused an intensive study in this area in order to improve its protocols. Thus, an improvement would make possible an increment in the target protein yield and the quality production as well as to establish more efficient host and plasmid for each target protein .
The most common hosts utilized in RPP are bacteria because of the capacity that they have to express almost any gen and the relative facility to modify and use their plasmids in order to produce the target protein .
However, it is widely known the number of problems that these hosts have when they produce a high amount of recombinant protein. Firstly, a frequent problem is the appearance of inclusion bodies which hinder a correct recovery of the target protein produced . Secondly, the host lysis event is the other common problem in RPP in bacteria. This undesirable happening in the production of recombinant proteins can be produced for several reasons. One of them is the high level synthesis of the mRNA and the target protein . Other reasons described are the accumulation of fragments of the recombinant protein because of the proteolysis . Finally, the main cause of the problems related to RPP is the accumulation of incorrectly folded intermediates of the recombinant protein. In E. coli this fact implies general stress responses .
In order to find a proper protocol for the protein S (PS) production in E. coli and know more about the RPP process, we conducted an experiment testing post-induction bacteria growth, production of target protein and plasmid retention. In this experiment the E. coli strain BL21*DE3 transformed with the plasmid pCV05 was used to express the PS fused with a C-terminal His tag (6xHis).
Materials and Methods
The E. coli strain BL21*DE3 transformed with the plasmid pCV05 (a derivative of pET21a plasmid) was utilized to conduct the experiment. This plasmid carries in its sequence the gene of PS fused with a C-terminal extra sequence which encodes a His tag (predicted molecular weight 60 KDa). A flask with 25 mL LB (10 g L-1 tryptone, 5 g L-1 yeast extract, 5 g L-1 NaCl) supplemented with 0.1 g L-1 carbenicillin was inoculated with a single colony of E. coli. Subsequently, this flask was incubated during 7 hours (25°C). The next step was to inoculate the 2.5-L fermenter (ΑG CH-4103 Bottmingen®), which contained 2 L LB supplemented with 0.5% (w/v) glucose and 0.1 g L-1 carbenicillin, with 25 mL from the flask previously inoculated and incubated during 8 hours (25°C). Immediately before inoculate the fermenter, a sample was taken in order to use it as a blank to measure the OD650nm of the following samples. The culture was grown at 25°C until an OD650 nm of nearly 0.6 when it was added the inducer of recombinant protein expression (IPTG). The culture was supplemented with 100 µM IPTG. After this step, the culture was grown at 25°C during 8 hours. The pH was controlled at 7 adding 5% (v/v) HCl and 1 M NH3, and 0.03% (v/v) final concentration of silicone antifoam was added in the fermenter due to prevent foaming during the last hours of the fermentation. During that period of 8 hours, the culture was fed and samples were taken at various times. One sample was picked just before the induction (0h) and the other samples were collected at 1h, 3h, 4h, 5h, 6h and 7h after the induction in order to check the bacteria growth. Besides, a pellet of bacteria was obtained from those samples collected to check the amount of target protein by a 15% SDS-PAGE gel subsequently stained with 0.2% (w/v) Coomassie Blue using the NEB Prestained Protein Marker, Broad Range (7-175 KDa)® as a marker. In addition, BugBuster Protein Extraction Reagent® protocol was conducted for additional 0h, 3h, 4h and 8h samples due to separate the insoluble and the soluble protein fractions from the bacteria and assess them by SDS-PAGE as described above. On the other hand, 0h, 3h and 7h samples were plated onto non selective nutrient agar (NA) and later replicated onto NA supplemented with 0.1 g L-1 carbenicillin in order to assess the plasmid retention of the bacteria.
Results
Inducer effect in Bacteria growth
It was conducted a measurement of OD650 nm of the culture during the 8h-period of growth. In order to conduct that assessment, samples were collected immediately before the induction (0h) and 1h, 3h, 4h, 5h, 6h, 7h after the induction with IPTG.
After the IPTG induction the culture grew slowly until time 3h. After that time, the growth rate was increased significantly until time 5h. However, at this time the culture presented a decrease in growth until time 7h. The growth dropped specially between time 5h and 6h (Fig. 1).
Plasmid retention
To assess how the E. coli strain BL21*DE3 retains the pCV05 plasmid after the IPTG induction, samples from the fermenter were collected at time 0h, 3h and 7h post-induction. Serial dilutions of these samples were plated onto non selective NA and subsequently replicated onto NA supplemented with carbenicillin. The relation between the colonies grown on NA plates and the colonies grown in NA supplemented with carbenicillin give the % of plasmid retention.
The plasmid retention was hardly altered during the period of 7 hours post-induction assessed. Unexpectedly, according to the results obtained, the plasmid retention 3 hours after the induction is higher than the rate at time 0h. Regarding the time 7h, the plasmid retention percentage is much lower than in time 0h and 3h (Fig. 2).
Target protein yield
In order to compare the target protein yield during the growth of the culture, samples were collected immediately before the induction (0h) and different times post-induction (1h, 2h, 3h, 4h and 5h). In addition, insoluble and soluble fractions for time 0h, 3h, 4h and 8h were obtained in order to know if the target protein was correctly folded (protein in soluble fraction) or incorrectly folded (insoluble fraction). These samples were assessed in a SDS-PAGE gel.
According to the SDS-PAGE profiles, although the PS::His band should be around 60 KDa, in this experiment the target band seems to be below the expected weight (Fig. 3).
Regarding the total protein assessment, this band shows a gradual increase in its intensity from time 3h to time 5h post-induction. However, at time 0h, 1h, 2h after induction there was not significant production of the recombinant protein (Fig. 3a). As for the soluble and insoluble fraction samples, the SDS-PAGE analysis revealed a higher intensity of the target protein band in the insoluble fraction than in the soluble fraction of the times 8h, 4h and 3h. At those times, the intensity of the recombinant protein band for insoluble fraction samples is almost impossible to appreciate. The same occurs for the soluble and insoluble fraction samples at time 0h (Fig. 3b). Overall, according to the results of the SDS-PAGE profiles, there was a significant recombinant protein production since the time 3h after the induction with IPTG. On the other hand, it seems to be that there was not a proper target protein folding because the recombinant protein could be found in the insoluble fraction rather than in the soluble fraction.
Discussion
The main cause of an incorrect folding of the recombinant protein when a high yield is conducted is well known. The accumulation of misfolded protein intermediates causes considerable stress in the host cell . A wide range of different strategies have been conducted in order to solve this problem . It has been described that the use of IPTG-inducible T7 RNA polymerase system in the BL21 strain to produce high concentrations of recombinant protein usually implies a high level of post-induction stress . The solution proposed by some researchers is select mutants which have lower expression rates of the recombinant protein . Other researchers have opted for limiting the concentration of inducer used . Nowadays, the establishment of general protocols and host for the different target proteins is still a challenge.
In this experiment, we have assessed the capacity of the E. coli strain BL21*DE3 to produce a high amount of PS fused with a C-terminal His tag as well as the bacteria growth during the process and the plasmid retention. According to the first part of the results, the bacteria growth rate changed during the 7 hours of fermentation. It is described that the high amount of recombinant protein in the cell causes stress response. This stress response implies that the growth rate of the culture turned into a negative rate (the number of cells in the culture decreases) . The data collected from other groups conducting the same experiment shows that this event has happened in 2 groups but the other 3 groups have a different growth pattern (Fig. 4). Technical issues may explain this incongruity between the different results obtained.
Regarding the plasmid retention, the data obtained in this experiment suggest that because of the stress suffered by the cells when the recombinant protein levels are higher, the cells tend to have a lower rate of plasmid retention. An explanation may be that the bacteria with the plasmid suffer a higher stress due to the induction by the IPTG, and thus, they have less chance to survive than the bacteria which accidentally do not have the plasmid. Therefore, the bacteria suffer a selective pressure which results in a plasmid loss and it is more obvious after several hours of growth. Analyzing the data obtained by other colleagues, it is supported that there is a relation between the time after the induction and the plasmid retention rates (Table 1).
As for the PS::His yield, the data shows that in the BL21*DE3 strain using pCV05 as a plasmid and with the conditions described before, this expression system needs 3 hours to start expressing the recombinant protein. After that time, it seems that the most part of the target protein is in the insoluble fraction. Regarding the total protein samples assessment, the relative amount of target protein produced by the cultures of the all groups was the same (Table 2). However, the recombinant protein is still in the insoluble fraction after the BugBuster Protein Extraction Reagent protocol. Probably, some of the proteins from the insoluble fraction were forming inclusion bodies . It is described that after the expression of the protein removing the His tag makes the recombinant protein more soluble and thus, it is possible to dissolve the inclusion bodies and recover a functional recombinant protein . Furthermore, the His tag allows an easier purification of the protein due to the affinity of this polypeptide for metal ions . In order to confirm those statements for PS, it would be necessary to conduct purification and a proteolysis process of the His tag in order to assess whether the efficiency of this expression system is cost-effective and the amount of protein obtained is enough.
The conclusion of these data is that this expression system for the PS yield could be a good and profitable system whether the His tag added finally allows to recover the recombinant protein from the inclusion bodies.
Acknowledgements
We are grateful to Dr Claire Vine, Dr Ian Cadby and Dr Jeff Cole for the excellent support given as well as the rest of the groups which conducted the same experiment because they have contributed to the experiment with very valuable data.
 

Microflora and Bacteria in Limburger Cheese Development

Limburger Cheese

Introduction

Limburger cheese gets its name from the country it originated in, Limburg, which is now divided amongst Germany, Belgium, and the Netherlands.1 Limburger cheese is made from pasteurized cow’s milk and has a 42% fat content.1 This cheese is typically more mild due to the milk that is used in the production process.1 The texture of this cheese can be characterized as creamy, crumbly, firm, and smooth.1 However, this cheese is best known for its highly stinky aroma resembling that of foot odor.1

Limburger cheese is rind washed. Due to the regular rind washings, the exterior of the cheese is covered with a thin pale, orange-brown rind.1 Starting out, the Limburger cheese is firm yet crumbly; this phase is very short before the cheese becomes chalky and soft.1 This chalky and soft characteristic takes place after about six weeks.1 The cheese becomes creamier and smoother at about two months.1 Then at three months, the cheese acquires its stinky aroma.1 The exterior portion of the cheese is the thin rind, while the interior of the cheese is a soft and yielding straw colored paste.1

Microflora

Initially, the microflora of bacterial smear surface-ripened cheeses appear to be very similar at the beginning of the ripening phase; however, the bacteria that becomes present at the end of the ripening stage is what sets the cheeses apart.2 The limburger cheese aroma is due to smear ripening with solutions of bacteria.1 “Early studies on the microflora of Limburger showed that when the pH of the cheese surface rises to 5.85, due to the growth of the yeasts, the growth of B. linens commences.2”

Yeast

Yeast can establish significant interactions with the bacteria present on the surface and lactic acid bacteria.2 Smear surface cheeses contain the most bacteria on the surface of the cheese such as the rind and play main roles in the final characteristics and attributes.2 Yeasts and molds initially dominate the surface post manufacture because they are acid tolerant and salt tolerant, but at the end of the ripening period, bacteria of the genera Brevibacterium, Arthrobacter, Micrococcus and Corynebacterium are the dominant microorganisms.2 Although the yeast play an important role in the smear ripening process, very small yeast cells are present once the cheese ages.4

Yeast also play an important role in the bacteria smear ripened cheeses. The yeasts interact with the molds and bacteria including on the surface and the lactic acid bacteria.2 The yeasts also help contribute to the flavor, texture, and aroma to the cheeses.2

The first thing to develop on the external part of the smear ripened cheeses is yeast.2 “Studies on the evolution of yeast populations on the surface of cheeses such as Limburger (El-Erian, 1972) have shown that they reach the highest number of 108–109 cfu/g of smear after about 7 days of ripening. After that time, a stable value of the yeast population, has a decrease in their number, as shown in Limburger cheese, have been reported. However, the general trend is the yeast domination during the early stages of ripening, followed by bacterial domination of the surface flora.2”

Bacteria

Clostridium licheniforme and Paraplectrum foetidum are two other bacteria that have been studied and found on the surface of the smear ripened Limburger cheese.4 After research by Weighmann he came to discover that the two prior bacteria were aiding in breaking down the lactic acid in the cheese and making the medium more alkaline.4 Weighmann also discovered that a red bacteria that would cover the smeared surface of the cheese, was anaerobically permitting the growth of P. foetidium.4 Weighmann also credited the aroma and flavor of the Limburger cheese to the bacterium P. foetidium.4

Chemical Compounds

Smear surface ripened cheeses happen to ripen faster and have more of an intense flavor profile than other cheeses.2 The reason for the fast ripening and the strong flavoring is due to poor syneresis properties such as, the method of curing involved which allowed the creation of favorable optimal development for the microbiological community.2

The two main microorganisms present in Limburger cheese is Streptococcus thermophillus and Lactobacillus delbrueckii subsp. lactis. These two microorganisms are responsible for providing distinct flavors and textural attributes.3 Limburger cheese is coagulated by rennet, then goes through an external bacterial ripening stage.3 Limburger is a semi soft cheese with a moisture content ranging from 39-50% with 50% being the maximum moisture content allowed in this specific cheese.3 This cheese is surface ripened by bacteria and yeast.3 Due to the salting method that takes place during the ripening of Limburger, salt tolerant microorganisms grow very well on the surface at this time.4

 

Background

There are 17 species of the Listeria genus; however, only Listeria monocytogenes is pathogenic to humans.6 L. monocytogenes is can be fatal to humans due to its widespread diffusion in the environment and food.6 Ready-to-eat foods have been traced as being the main vehicle for transmission of Listeria through contamination somewhere in the food chain.9

L. monocytogenes is also a dangerous pathogen, due to its ability to withstand and grow in low temperature environments and high salt concentrations.6  “Compounds used to inhibit growth of this pathogen include organic acids, fatty acids, antioxidants, sodium nitrite, smoke, spices and herbs.9” Another reason L. monocytogenes is a dangerous pathogen is because “some strains are able to survive for long periods of time under adverse environmental conditions and persist in niches in food processing equipment, and associated drains, walls and ceilings.9” However, most large outbreaks are contracted through errors in food processing plants.9

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 There have been multiple different studies done showing if L. monocytogenes can survive through the process of pasteurization or not. “An early study by Bearns and Girard showed the L. monocytogenes may be able to survive pasteurization if present in fresh milk at concentrains of more than 5 x 104 organisms / ml.Conversely, Bradshaw and collaboratorsfound that L. monocytogenes could not withstand pasteurizing temperatures.7” The only way the pathogen would be able to withstand the pasteurization would be if it resides within leukocytes.7 Another way L. monocytogenes can withstand pasteurization is through its ability to form inside of biofilms.9 Overall, L. monocytogenes is more likely to be contracted through the consumption of raw milk.6

“L. monocytogenes infects normally sterile parts of the body such as the liver, spleen, cerebral spinal fluid and blood.9” Most cases of this pathogen result in being hospitlaized.9 For healthy adults symptoms mainly include diarrhea and fever, for pregnant women symptoms include fever, diarrhea, abortion or still-birth, and symptoms in newborns include sepsis, pneumonia or meningitis.9

Listeria monocytogenes in Semi-Hard Cheeses

The growth potential of Listeria monocytogenes on semi-hard cheeses, such as Limburger, was generally lower than the other types of cheeses.5 The range of L. monocytogenes growth potential ranged from 0.1 to 1.4 log units at 7 °C and from 0.0 to 3.0 log units at 14 °C.5 “Overall, increased outgrowth of L. monocytogenes was noted when inoculation was performed on the cheese slicing surface compared to the cheese rind.5” There is a relatively large variation in the presence of L. monocytogenes in different cheeses.5 The storage temperature and the cheeses type play a role in the growth potential of L. monocytogenes.5

Although, semi-hard cheeses have a lower growth potential of obtaining L. monocytogenes, it has the possibility to grow in numbers through post-processing contamination, storage, or handling of the cheese up to 14 days notwithstanding the presence of high numbers of indigenous lactic acid bacteria in these cheeses.5 It has been tested that the most frequent cases of cheese-related outbreaks with L. monocytogenes occurs in the post-pasteurization contamination.8 In Europe, human foodborne infections of L. monocytogenes were identified as the vehicle for the soft and semi-soft cheeses.5

“It was also found that semi-hard cheeses will not support growth, but only enable survival of L. monocytogenes. The growth potential in soft and semi-soft cheeses, on the other hand, is noted to be substantially higher than in semi-hard cheeses. Thus, it can be concluded that soft and semi-soft cheeses present indeed a higher concern with regard to listeriosis.5”

L. monocytogenes was the most resistant of all bacteria tested.8 It survived in the interior of the semi-hard cheese for more than 90 days of ripening.8 L. monocytogenes grew profusely on the surface of the semi-hard cheeses.8 Even after the intense brining and ripening at elevated temperatures for at least 2 months L. rnonocytogenes, at the time of commercial ripeness was still present.8 The optimal water activity in the milk, curd, and cheese for the pathogen, L. monocytogenes is 0.998-0.999.10

Smeared ripened cheeses made from raw cow’s milk have also been identified as a vehicle for L. monocytogenes.5 This is where most of L. monocytogenes begins and stays in the pre-production of cheese because this pathogen can get into the biofilms and stay there for longer periods of time.5 Dairy farms is where this pathogen can be introduced if proper hygiene and sanitation is not taken place, but this pathogen can also have an outbreak in post-production or further distribution.5  However, “the epidemiology of cheese-related outbreaks demonstrates that surface-ripened soft cheeses generally represent a greater risk for the transmission of pathogens than do other cheeses.8”

“L. monocytogenes can occur during storage, display or slicing when the bacterium colonizes the environment, equipment, utensils and crates.It was also expected that more L. monocytogenes growth would occur on the cheese slicing surface than on the cheese rind.5” The growth of L. monocytogenes is not supported by hard cheeses, however, hard cheeses may support the survival of L. monocytogenes.5 “Moreover, some studies demonstrated that L. monocytogenes will die during ripening of hard cheeses.5”

There is a difference in the amount of L. monocytogenes present depending on what type of milk is used. Along with what kind of milk the cheese is being made with, the type of distribution can also have an affect of L. monocytogenes growth this includes a local market, small shop, or big retail outlet.5 One of the last affects of the presence of L. monocytogenes is the selection of regions where the samples of the milk is taken from.5 “It could not be observed from the challenge tests that growth of L. monocytogenes in pasteurized cheeses is significantly higher than growth in raw milk cheeses.5” It was proven; however, that cheese made from raw milk had a slower growth rate of L. monocytogenes compared to that of pasteurized milk.5 “Mathew and Ryser (2002) observed that heat-injured cells of L. monocytogenes were recovered with higher rates in heat-treated or pasteurised milk than in raw milk.10”

“This difference in growth potential of L. monocytogenes may be explained by the presence of the lactoperoxidase enzyme in raw milk cheese which has bacteriostatic properties in milk and milk-based products. In this study, it was noticed that pasteurized milk cheeses have lower contamination levels of E. coli than raw milk cheeses. This is due to the heat treatment used during processing. Therefore, more bacterial competition is expected to be present in raw milk cheeses which may be as well an explanation of the lower growth potential of L. monocytogenes in these cheeses  Although the prevalence on pasteurized cheese may be lower, if there are opportunities for growth of the pathogen, higher numbers of L. monocytogenes may be obtained in pasteurized cheeses, making raw milk cheeses and pasteurized cheeses equally important in terms of at risk products for listeriosis.5”

L. monocytogenes may not have a large presence in cheese, but it definitely has some sort of presence. “L. monocytogenes had not yet exceeded numbers of 100 CFU/g. As mentioned before, the presence of low numbers of L. monocytogenes in cheese is not an infrequent event.5” Once this number rises above a certain point, is when this pathogen becomes dangerous to humans and consumption. 

Results

A study of Gould, Mungai, and Behravesh (2014) where 90 outbreaks in the United States attributed to cheese were analyzed.5 The study showed that 42% of the outbreaks were due to cheese made with unpasteurized milk and 49% due to cheese made with pasteurized milk.5 Only 12 of these outbreaks were caused by L. monocytogenes.5Only 4 out of these 12 were involved with unpasteurized milk cheese.5 The remaining 8 out of the 12 outbreaks were involved with pasteurized milk cheese.5 Overall, soft raw milk cheese holds the greatest risk for survival and growth of L. monocytogenes although the growth potential will ultimately depend upon the actual storage temperature.5

More results from this study demonstrate that “higher growth of L. monocytogenes is obtained on a sliced surface of the cheese than on the cheese rind. It was also shown that in both raw and pasteurized semi-soft washed-rind milk cheeses, the L. monocytogenes population increased as the temperature increased.5” In specific cheese types, however, in neither the inter-batch nor intra-batch variability was recorded having consistent behavior with L. monocytogenes in fermented dairy products such as cheese and milk.5

Pasteurized milk was found to favor the growth of L. monocytogenes during cheese making; moreover, there was no growth of L. monocytogenes during cheese making with raw milk.10 The ripening period of the cheeses revealed that the cheeses made with pasteurized milk had a decrease in numbers of L. monocytogenes present on the rind.10 In contrast, the ripening period of the cheeses made with raw milk revealed L. monocytgenes grew in raw milk cheese, but was inactivated in the pasteurized milk cheese.10 “In this study, we observed that L. monocytogenes grew during the manufacture of pasteurized milk cheese, whereas during the manufacture of raw milk cheese, no growth of the pathogen occurred.10 Morgan et al. (2001) also found that L. monocytogenes survived the ripening of soft lactic cheese made with raw milk, while Margolles et al. (1997) found that L. monocytogenes was inactivated during the manufacture, ripening and storage of an artisanal acid-coagulated cheese made with pasteurized milk.10” The main difference between the raw milk and the pasteurized milk was pH and lactic acid concentration.10

To prevent the spread and contamination of L. monocytogenes, one must ensure the safety of the post-processing contamination and be aware that contamination can occur from the farmer, to retailer, to salesmen. Being aware and taking proper hygiene precautions can limit the risk of listeriosis.5

Listeria monocytogenes in Raw Bovine Milk

L. monocytogenes can originate inside of an infected cow’s ruminant mastitis.6 This specific case was found in a herd in Italy.6 Health inspectors went to the dairy farm that the infected bovine milk came from to test to see which cow or cows were infected with L. monocytogenes.6 To test for this, each cow was milked and their milk was tested for the presence of this pathogen. One cow in the herd was found to be producing a persistent excretion of this pathogen.6 L. monocytogenes can stay present in the milk for months after a contamination and does not have any antimicrobial treatment.6
 

Techniques for Identifying Unknown Bacteria

Section I: Introduction

  Microorganism surrounds the environment which makes them part of our health and well-being, but sometime, they can become harmful and the ability to correctly identify the harmful species is extremely useful in applied microbiology. New technologies and discoveries are happening everyday, but basic microbiology lab techniques and concepts are needed to perform an accurate bacteria species identification and understanding how these techniques work is also important. A few of these concepts include the aseptic method, microscopic examination, selective and differential media and the importance of having a positive and negative control.

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The aseptic technique is the most basic and crucial method of culture transfer in order to have the most accurate test results as possible. This technique consist of a transfer method from various types of media by using a microincinerator sterilizer device that produces high heat which reduces the risk of contamination while performing a culture transfer. An inoculating loop or needle is the instrument used to perform the transfer and has to be inserted in the microincinerator before and after each transfer of culture. The aseptic method is crucial in order to isolate pure cultures, as the inoculating loop is sterilized between each step of the culture transfer which eliminates contaminants and leads to the growth of pure culture (1). Contaminants that are present in growth culture can lead to misinterpretation of results and ultimately sabotage the identification of the unknown bacteria, for this reason, using the aseptic method is the most crucial method of culture transfer.

Microscopic examination is another crucial technique that needs to be mastered as it allows to observe cultures with greater magnification and resolution. This technology permits to distinguish the morphology of the unknown culture and ultimately helps eliminating bacteria that do not fit the characteristics observed via the microscope (2). The microscope has evolved from when it was first introduced by Van Leewenhoek whom was the first scientist to accurately describe bacteria morphology (2). Nowdays, there are several kinds of microscopes such as electron microscope or light microscope, the latter has several subcategories where the bright field microscope is the most popular in a typical microbiology laboratory setting. Bright field microscope allows to see specimen with light coming from underneath and makes it darker than its surroundings (2). Using the differential Gram staining method prior to microscopic observation is often done to give additional information such as classification of the bacteria as Gram positive or Gram negative (1). Knowing the classification also sets the stage for which subsequent tests will be performed in order to determine the identity of the unknown bacteria.

To perform a series of tests, the unknown culture needs to grow on an appropriate media in order to receive the right nutrients and ensure its growth and survival, this is where the use of differential and selective media are employed.  A selective medium contains all the basic nutrients needed, but also has a few components added that inhibits the growth of undesired bacteria (l). A differential medium allows to distinguish between species that are capable of performing certain biochemical processes, which further assists in the identification process of the unknown culture (1) It is important to note that some media can be classified as both selective and differential.

While performing the tests, keeping track of the positive and negative controls is crucial because it is used as a baseline to compare the unknown test results. A positive control for a test means the bacteria performs the reaction tested and a negative control means the bacteria is not capable of performing the reaction tested. Having these controls as a guideline is essential as it eliminates confusion in test results obtained from the unknow culture, and consequently, narrows down possible errors while identifying the unknown culture.

With all the possible species the bacteria domain can have, a sytematic bacterial classification can be challenging to come up with. A useful tool for bacterial classification is the The bacterial species concept which implies that bacteria with a common ancestor share similar physical traits as well as sharing similar genes (3). The biological concept takes it a step further where it implies that biological species should be able to reproduce by interbreeding and cannot breed with a non-self breed due reproduction isolation mechanisms (3). The biological concept is accepted for Eukaryotyic species classification, because genes are transferred from parents to offsrings, but in the case of bacteria this process is not observed, instead bacteria transfer genes via Horizontal Gene Transfer (1). The bacterial species concept is more suited to categorize bacteria species as the concept bases itself on DNA-DNA hybridization, 16S rRNA sequence data and horizontal gene transfer.

 DNA-DNA hybridization is among pioneered technique to measure how close the genomes of two bacteria are (4). This technique calls for precise manipulation such as denaturation and blending the two genome to see how much of it anneals together, the threshold of 70% DNA-DNA hybridization between is used to indicate that the two genomes are closely related (4). Althought the DNA-DNA hybridization is a good indicator of bacterial relatedness, it is not the most precise, because bacteria species can show a wide range of genome variety (3). To employ a more recent technology, scientits can use the 16S rRNA sequence comparison in addition to DNA-DNA hybridization method. Ribosomal RNA is highly conserved so it only makes sense that it evolves at a slow pace, which limits the genome variety issue when trying to find out bacterial relatedness (3). The 16S rRNA sequence is also easier to access and updated as it is a computerized database, but it is not infallible to the genome variability, in part due to bacterial horizontal gene transfer. Horizontal gene transfer as mentioned above, is the way bacteria exchange genome information and unlike Eukaryote that use lateral gene transfer, bacteria do not transfer genes from parents to offsprings (1). Horizontal gene transfer can be done via transformation, conjugation or transduciton, but regardless of the mechanism, the end result is the same. When bacterial transfer genes from one bacterium to another, it ultimately increases genomic variabiltiy. Despite all the effort mentioned above to compare bacterial genome whether it is through DNA-DNA hybridization or 16S rRNA sequencing, classification of bacteria remains a big challenge. In order to overcome these challenges, novel method called genomic-phylogenetic species concept (GPSC) is used. This concept uses both genomic and phylogenic analysis are combined to give a better outcome (5). GPSC is efficient and the most promessing because not only it analyzes DNA, RNA and protein, but it also looks at the phylogeny of the bacteria, which gives insight on the common ancestor and can relate species together even if they are not in the same location (staley)

Section II: Materials and Methods

 The unknown #118 was given to which a series of chosen tests had to be performed in order to identify the unknown bacteria. The unknown #118 came in tryptic soy broth (TSB) to which a serial dilution followed by an aseptic transfer using inoculating loop onto a Tryptic soy agar plate (TSA) was performed (1). Streaking method onto the TSA plate and incubation for a period of 24-48h at 25ºC was necessary for isolation of colonies (1). Two types of colonies were successfully isolated on the TSA plate, one large and the other small. The large colony was white, round, smooth, shiny, and the small colony was punctiform, also white, round, shiny and smooth.

Microscopic Examination and Gram Staining

Both colonies were subjected to Gram staining followed by microscopic examination under 1000X oil immersion using a bright-filed microscope. Gram staining was performed using crystal violet (1min) primary stain, iodine (1min) mordant, 95% ethanol (12-15sec) decolorizing and dehydrating agent followed by immediate distilled water rinse and  finally, safranin (1min) couterstain (1). Gram positive bacteria retains the primary stain crystal violet even after the decolorizing agent because of its thick peptidoglycan membrane (20-80nm) characteristic and can therefore be observed as dark purple under the microscope (l). On the other hand, Gram negative bacteria has a thin cell wall (2-7nm), the decolorizing agent is able to strip away the primary stain and would appear clear under the microscope, the use of a counterstain such as safranin is necessary to stain it pink for easier microscope observation (1). When the large colony was observed under the microscope, it turned out to be Gram negative with rod shape bacteria and the small colony was Gram positive with  short rod shape bacteria as well. The results were compared to the Gram negative control Escherichia coli and the Gram positive control Staphylococcus epidermidis.

Differential and Selective Media

 Mannitol Salt Agar (MSA) is a differential and selective media that contains high salt concentration (7.5% NaCl) that selects for bacteria capable of sustaining such environement (1). The media also has methyl red as ph indicator that turns yellow under acidic condition which indicates sugar is fermented by bacteria (l). The unkonwn #118 was aseptically transferred from TSB onto MSA plate via streaking method, followed by an incubation of  24-48hrs at 25º (1). Positive control for MSA test is Staphylococcus aureus (BSL-2) showed growth and yellow color on plate due to lowering of pH caused by acid build up from sugar fermentation, and  negative control for MSA test Staphylococcus showed no growth on plate.

 Maconkey (MC) is another differential and selective media that contains lactose, bile salts which inhibits Gram positive bacteria growth (1). The media contains neutral red ph indicator which changes color to pink when lactose fermenter is present due to acid release (1). The unkonwn #118 was aseptically transferred from TSB onto MC plate via streaking method, followed by an incubation of  24-48hrs at 35º (1). Positive control for MC test Escherichia coli showed pink growth due to acid build up caused by lactose fermentation, and negative control Pseudonoms fluorenscens showed colorless growth.

Catalase Test

Catalase test is used to determine if the bacteria is able to break down hydrogen peroxide (H2O2) into oxygen gas (O2) via calase enzyme. A loopful of the unknown #118 was deposited on a clean slide to which a drop of hydrogen peroxide (3%) was added on top (1). If bubbles are present, this would indicate that the catalase enzyme is present and bacteria is capable of breaking down H2O2 as observed in the positive control test Micrococcus, negative control Enterococcus showed no apparent bubbles.

Oxidase Test

 Oxidase test was used to determine if the bacteria has cytochrome c in the elecrtron transport chain (1). This test was perfomed by using an oxiswab imbedded with chromogenic redusing agent (TMPD), the cotton was swabbed inside the TSA plate containing the unknown #118, where a good amount of colony growth was present (1). If cytochrome c was present in the electron transport chian, the TMPD gets oxidized and turned deep purple/blue color just similar to the positive control Pseudonomas fluorenscens. On the other hand, negative control Escherichia coli did not show a change of color due to absence of cytochrome c in the electron transport chain.

Motility test

Motility test was used to give additional morphological information regarding the unknown #118 such as if the bacteria is motile or non-motile. The motility test was performed using an inoculating needle to aseptically transfer form unknown TSB onto a soft agar (0.4%) deep tube (1). The inoculating needle was inserted in the middle  of the soft agar (0.4%) in a precise linear fashion, without breaking the agar, and be removed before hitting the bottom of the deep tube in the same linear fashion. The tube was incubated for 48h at 35ºC (1). The use of soft agar (0.4%) is important in this test, because the bacteria needs to be able to move around in case it is motile, otherwise the bacteria would not be able to move and give a false negative test result. It is also important to not break the agar while performing the test, because this can lead to cloudiness and could be interpreted as a false positive result. The positive control  Proteus mobilis had a cloudy appearance due to the bacteria moving around the agar, and negative control Staphylococcus epidermidis (BSL-2)  was clear with only the stab print clearly visible in the tube.

Aerotolerance test

 Aerotolerance test was performed to determine bacteria oxygen requirment. There are different degree to which bacteria require oxygen, aerotolerance test can differentiate between obligate aerobes, facultative anaerobes and obligate anaerobes (1). This test was performed by using an inoculating needle to aseptically transfer from TSB onto agar deep stab tube followed by an incubation period of 48hrs at 35ºC (1). The inoculating needle was carefully inserted in a linear fashion, and removed before hitting the bottom of the tube via the same line of entry. While inserting the needle, it was important to not crack the agar, because it would lead to false test results (1). The controls displayed three different kinds of results: obligate aerobe Micrococcus luteus only showed growths at the very top of the agar, facultative anaerobe Echerichia coli had growth throughout the tube but mosty on top, and obligate anaerobe Clostridium sporogenes had growth toward the bottom of the tube.

Litmus test

The litmus test was performed to find out if the unknown #118 was part of the order Lactobacillales, which are lactic-acid fermenters or to differentiate the members of the family Enterbacteriaceae (l).  An inoculating loop was used to transfer from TSB onto litmus milk test tube and was incubated for 7-14 days at 35ºC (lab). Litmus milk test tube contained skim milk, oxidation-reduciton indicator, azolitmin pH indicator, and Na2SO3 (1). The skim milk consited of nutrients such as casein and whey proteins (1). A different array of results was observed from the controls, where the pH indicator changed color to red to indicate acidic conditions and blue for basic conditions (1). The redox indicator appeared white or colorless in reduced conditions and purple in oxidized condition (1). Uninoculated control had purple appearance, positive control for lactose fermenters were red/pink because of build up of acid and negative control were purple/blue which indicates alkaline reaction. Positive controls: Escherichia coli entire tube was pink, Lactococcus lactis had a pink acid clot and some white indicating litmus reduction. Negative controls: Bacillus subtilis had light purple, white, dark blue, Pseudomonas fluorenscens had some white at the bottom and purple.

Citrate test

 This test was used to determine if the unknown #118 was able to use citrate as the sole carbon source. There are a few members of the Enterobacteriaceae family that  are capable of fermenting citrate, and they can be differentiates by using the citrate test (1). The test was performed by using an inoculating needle to aseptically transfer from TSB onto Simmons’ citrate slant. The needle stabbed the end of the slant about 5mm deep, then was dragged out of the tube via a zig-zag motion followed by an incubation time of 48hrs at 35º (1). Uninoculated Simmons’ citrate slant are initially green in color and turn blue if citrate fermentation takes place. The positive control Klebsiella mobilis was green after incubation while negative control Escherichia coli remained green.

Methyl red/ Voges Proskauer Tests

 Methyl red (MR) and Voges Proskauer (VP) tests were useful to determine if unknown #118 was capable of fermenting mixed acid or butanediol after it was depleted of all glucose (1). Methyl red test was used to detect acidic end products, where as Voges Proskauer was used to detect neutral end products (1). This test was performed by using an inoculating loop and aseptically transfer from TSB onto MRVP broth tube followed by an incubation time of 48hrs at 35ºC (1). After the incubation time, the mixture was separated in equal amounts into two new test tubes, one designated MR and the other VP. In the MR tube, 10 drops of methyl red were added and set aside, where as 5 drops of KOH and 15 drops of alpha-naphtol were added in the VP tube followed by vigourous mixing (1). Positive control for MR Escherichia coli, a mixed acid fermenter, was a red color and negative contol for MR Klebsiella mobilis was yellow/clear in color. Positive control for VP Klebsiella mobilis, a butanediol fermenter,turned into a red color and negative control Escherichia coli had a yellow/clear color.

References

 

Malwane S, Malwane SD. 2018. A Laboratory Manual Microbiology 12th ed. Morton Publishing Company, Englewood, CO.

 

Madigan MT, Martinko JM, Stahl DA, Clark DP. 2012. Brock Biology of Microorganisms 13th ed. Pearson Benjamin Cummings, San Francisco, CA.

Riley MA, Lizotte-Waniewski M. 2009. Population Genomics and the Bacterial Species Concept. Horizontal Gene Transfer Methods in Molecular Biology 532:367–377.DOI: 10.1007/978-1-60327-853-9_21

Goris J, Klappenbach JA, Vandamme P, Coenye T, Konstantinidis KT, Tiedje JM. 2007. DNA–DNA hybridization values and their relationship to whole-genome sequence similarities. International Journal of Systematic and Evolutionary Microbiology 57:81–91.DOI: 10.1099/ijs.0.64483-0

Staley JT. 2006. The bacterial species dilemma and the genomic-phylogenetic species concept. Philosophical Transactions of the Royal Society B: Biological Sciences 361:1899–1909.DOI: 10.1098/rstb.2006.1914

 

Identification of Unknown Bacteria with DNA Sequencing

INTRODUCTION

 Bacteria are microscopic, single-celled organisms that can be found in all kinds of environment, that includes inside and outside of other organisms. They are the most diverse and successful group out of all prokaryotic organisms. The goal of this lab is to identify the unknown bacteria by traditional cellular and biochemical tests and by DNA sequence-based methods of species identification.

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 The first step of identifying the unknown bacteria is to determine the shape, size, color, and other characteristics of it. These characteristics help us determine if the colonies are bacteria, yeast, or mold. Bacteria will appear flatter and liquidity compared to yeast which will generally be puffier. Mold will look fuzzy and has a volcano-like elevated center. Bacterial colonies have multiple shapes such as circular, rhizoid, irregular, or filamentous. The colonies can also differentiate with different edges such as entire, undulate. Lobate, filamentous, or curl. Bacterial cells can vary in shape. For example, round (cocci), rod-like (bacilli), or helical (spirochete).

 The next step of identifying the unknown bacteria is to determine if enzyme catalase is present in the bacteria or not. Catalase is an enzyme that is produced by microorganisms that live in oxygenated environments that neutralize the bactericidal effects of H2O2. The catalase will break down the H2O2 into oxygen and Water. In order to find out if the bacteria can produce catalase enzyme, a small sample of the bacteria will be mixed with 3% hydrogen peroxide and observed for bubbles.

 An oxidase test will also help identify the unknown bacteria. The oxidase test is used to reveal whether the bacteria can produce cytochrome c oxidase, an enzyme for the bacterial electron transport chain. Which means bacteria that test positive for the oxidase test are aerobic, which means it can use oxygen a terminal electron acceptor in respiration. Bacteria that test negative either cannot use oxygen as an electron acceptor or utilize a different cytochrome to transfer electrons to oxygen. For this test, we will be observing for color change on the oxidase slide.

 Then a Mannitol Salt Agar (MSA) test is performed. This test is used to test for salt tolerant. Because MSA contains a high concentration of sodium chloride, only the salt-tolerant species can grow on it. Non-salt tolerance species cannot survive to reproduce on this medium. And the fermenting species would change the color of the medium from red to yellow or orange.

 Next, we determined whether the bacterial sample was Gram-Positive or Gram-Negative. Gram-Positive bacteria have cell walls composed of thick layers of peptidoglycan while Gram-Negative have cell walls with a thin layer of peptidoglycan. With that, we started culturing our bacteria on three different mediums: EMB-Lactase, PEA, and vancomycin. Gram-Positive bacteria will grow on PEA medium only. While the Gram-Negative bacteria will grow on EMB-Lactose and vancomycin medium. The potassium hydroxide test (KOH string test) is another test that we did to determine if the bacterial sample was Gram-Positive or Gram-Negative. The KOH will dissolve the thin layer of peptidoglycan of the cell walls on the Gram-Negative bacteria, but it has no effect on the Gram-positive cell walls. The KOH will dissolve the cell walls and the content of the bacteria including the DNA will be released. Because of the DNA the solution will become viscous and it will stick to the toothpick creating a “string”.

 To perform the last test, which is gel electrophoresis, we used PCR to amplified DNA from our unknown bacteria. The 16S Ribosome gene will be isolated and amplified to provide us with the template for the DNA sequence used to help us identify the unknown bacteria. Once the DNA is sequenced it was edited in Chromas and ran through BLAST, a database that has millions of gene sequences, to be aligned with the best possible match allowing us to identify the unknown bacteria.

MATERIALS AND METHOD

Making a Live Culture

 To make the live culture, we scraped a colony of the unknown bacteria and transfer it to a PCR tube containing 295 μL of sterile water. Then we vortex the tube for 2-3 seconds to uniformly distribute the cells in the water (Holbrook and Leicht, 2019).

Setting up PCR Tubes

 Once the live culture has been created, we set up the PCR tube. We made two PCR tube; one with bacterial DNA and the other for control which contains no DNA and both containing 25 μL of 2X PCR Master Mix. We added 5 μL of live bacterial culture and 20 μL of 16S rRNA Primer Mix to the tube with bacterial DNA. In our control tube, we added 5 μL of sterile water and 20 μL of 16S rRNA Primer Mix. Then both tubes were briefly centrifuged in the microcentrifuge. Once mixed, the DNA was amplified by the following procedure (Holbrook and Leicht, 2019):

1X: 94oC, 3 min

30X: 94oC, 30 sec; 50oC, 30 sec; 72oC, 45 sec

1X: 72oC, 5 min

Purification of PCR Products

 After the DNA amplification is done, the content in the tube with bacterial DNA was transfer to a new 1.5 mL microcentrifuge tube. Next 250 μL of Buffer BB was added to the PCR sample. Then the PCR sample was transferred to a spin column in a collection tube and centrifuge for 30 seconds at room temperature in the Eppendorf5430. Next, the flow through was discarded and placed back into the collection tube. 200 μL Buffer WB was added to the spin column and centrifuge for 30 seconds in the Eppendorf5430. The flow through was discarded again. Then the process of adding WB Buffer, centrifuge, and discarding was repeated. Then the spin column was transferred to a 1.5 mL microcentrifuge tube. Next, 25 μL Buffer EB was added to the center of the membrane of the spin column and left to stand for 1 minute. After 1 minute, it is centrifuge for 30 seconds in the Eppendorf5430. Then the spin column was discarded and the 1.5 collection tube that contains the DNA will be used for gel electrophoresis and sequencing (Holbrook and Leicht, 2019).

Catalase Test

 To conduct the catalase test, we obtained a microscope slide and place it in an empty Petri dish. Then the unknown bacterial were collected and smeared onto the microscope slide. We added one drop of 3% hydrogen peroxide onto the smeared bacterial and watched for bubble formation (Holbrook and Leicht, 2019).

Oxidase Test

 First, we obtained a dry oxidase slide with four test areas for an oxidase-positive control, an oxidase negative control, and for two unknown bacteria. We collected each bacterial sample to spread onto the corresponding areas of the oxidase slide, then leaving the oxidase slide to incubate at room temperature for at least 20 seconds. After 20 seconds, an oxidase-positive bacterium should exhibit a color change (Holbrook and Leicht, 2019).

MSA Test

 For the MSA test first, we added the unknown bacteria to a tube containing sterile water. Then, using a micropipette we added 50-75 μL of the liquid bacterial culture and 5-10 sterile glass beads to MSA containing Petri dish. The Petri dish was shaken up and removed once the liquid bacterial culture is evenly distributed. They were incubated for 2 days at 37oC before we could observe the growth on the dish (Holbrook and Leicht, 2019).

Plating on Test Media

 A liquid culture containing unknown bacterium and 300 μL of sterile water was added to EMB-lactose, PEA, and vancomycin containing agar. The 5-10 sterile beads were also added and remove once the liquid culture is distributed evenly. The Petri dishes were incubated for 1-2 days at 37oC before we could observe the growth on the dish (Holbrook and Leicht, 2019).

 

 

KOH String Test

 50 μL of 3% KOH were added to a microscope slide with the unknown bacteria. The solution was stirred for about 60 seconds. Then using a sterile toothpick, we test to see if the solution has a “string” appearance (Holbrook and Leicht, 2019).

Preparing the Gel for Analysis of Purified PCR Product

 To prepare the 40ml of 1.5% agarose solution we added 0.6g of agarose and 40ml of 1X TBE buffer into a glass flask. Then we microwave the solution for 45-60 seconds until the solution boiled. While the gel is cooling, we set up the gel tray. After the flask has cooled to the point where it is warm but not hot, ethidium bromide was added to the flask. Then we gently swirled the flask to mix the solution. Once mixed we poured the liquid into the gel tray and leave it to set completely. When the gel has solidified, we removed the comb and placed the tray with the gel onto the electrophoresis chamber and filled the chamber with 1X TBE buffer until the gel completely covered with buffer (Holbrook and Leicht, 2019).

Loading and Running the Gel

 First, we added 6 μL of purified 16S rRNA PCR products and 4 μL Loading Dye into the microcentrifuge tube with the bacterial sample. As for the control microcentrifuge tube, we added 12 μL of the control PCR reaction and 4 μL of the Loading Dye. Then both tubes were spin for about 10 seconds in the mini centrifuge. Next, using the P20 micropipette we load 10 μL of bacterial DNA into wells 2, 5, and 7 (three different groups shared the gel, so each well had a different bacterial DNA), 10 μL of the controls were added to wells 3, 6, and 8, 10 μL of DNA Size Stand were added to well 4, and leaving well 1 empty. The lid was placed onto the electrophoresis apparatus and was set at a high voltage. Then we ran the gel until the blue dye moved about ¾ of the length of the gel. Once the gel had finished running, it was removed and photographed with the Fotodyne UV illuminator and camera (Holbrook and Leicht, 2019).

DNA Sequencing Reaction

 To create the dilution, we mixed together 2 μL of purified PCR product and 6 μL of sterile water. Then to get to the concentration of 3-10 ng of PCR DNA, we used 4 μL of the dilution and 6 μL of Big Dye mix. The tube was mixed and placed into the thermocycler. The following program was used (Holbrook and Leicht, 2019):

1X: 96oC, 1 min

30X: 96oC, 10 sec; 50oC, 5 sec; 60oC, 2 min

Hold: 4oC

Chromas and BLAST Database

 Once the DNA is sequenced, we used Chromas to analyze it. The poorly read ends were trimmed out, and the “N” basses are changed to whatever base had a peek at that location. Once edited, the sequence is copy and pastes into the BLAST database to determine the species of the bacteria (Holbrook and Leicht, 2019).

RESULTS

Cell Morphology

 When observing our petri dish containing the unknown bacteria, it was slightly raised with a yellow tint color throughout the colonies. All the colonies were circular with a smooth and glistening surface. When observing our live culture under a microscope, we observed the bacteria shape to be rod-like.

Catalase/ Oxidase Tests

Figure 1: Results of Catalase and Oxidase Tests.

When we performed the catalase test with our unknown bacteria, the hydrogen peroxide reacted strongly with the bacteria. There was an immediate formation of bubbles which was created by the conversion of H2O2 to O2 and H2O. As for the Oxidase test, the bacteria reacted strongly on the oxidase slide. When the bacteria were smeared onto the slide it turned blue indicating that the bacteria possess cytochrome c oxidase.

MSA Test

Test

Results

MSA Test

No growth

Figure 2: Results of the MSA test.

 The growth of the unknown bacteria was observed when it was placed on MSA. No growth was observed which means the unknown bacteria is a non-salt tolerant species.

 

 

Growth on Different Medium

Growth on Selective Medium

Forms a String KOH

Vancomycin

EMB-lactose

PEA

Amount of growth

+

N/A

Colony color

Yellow

N/A

N/A

N/A

Gram-positive or

Gram-Negative

Gram-Positive

Gram- Negative 

Gram-Negative

Figure 3: Classification of Unknown Bacteria as Gram-Positive or Gram-Negative.

Figure 4: Plate growth from 3 media. Vancomycin (Left), EMB-lactose, and PEA (right).

 The unknown bacteria were observed for growth on here different media; Vancomycin, EMB-lactose, and PEA. Only Vancomycin showed growth of a yellow color colony which indicates that it is a Gram-Negative Bacteria. Since there was no growth in EMB-lactose media, it indicated that the unknown bacteria are Gram-Positive. There was also no growth shown on the PEA media which means that the unknown bacteria are Gram-Negative. We also conducted a string test to help us determine if our bacteria are Gram-Positive or Gram-Negative. The string test indicated that our unknown bacteria is Gram-Negative.

Gel Electrophoresis

Figured 5: Gel electrophoresis of unknown bacterial DNA

1  B2  C3  SS B5 C6  B7 C8 88

 

 

 

 Figure 5 is the results of the gel electrophoresis of the unknown bacteria.

We can see all three lanes that have bacterial DNA. The band in lane B2 is half as bright compared to the size standard band. As for band in lane B5 and B7, it is as bright as the size standard band. The negative control lane (C3, C6, and C8) did not have any band which means there was no contamination.

Chromas and BLAST Database

Figure 6: The edited chromatogram from sequenced DNA.

Figure 7: Top 5 Match from the BLAST database for unknown bacteria

Figure 8: Best nucleotide alignment of Unknown Bacteria DNA sequence and the closest match species.

With the sequenced DNA, we were able to edit it and submit it to the BLAST database. With this, we found out that our bacteria stand was Aeromonas shown in figure 7.

Discussion

 The unknown bacterial DNA sequence of 16S rRNA gene matched up with the sequence of Aeromonas. Aeromonas is rod-shaped bacteria. Their colonies are smooth, convex, rounded and are tan/buff-colored. It is also Gram-Positive, Oxidase positive, and Catalase Positive. Aeromonas can exist in both aerobic and anaerobic environment. Aeromonas is also salt tolerance but up to a certain concentration.

 The characteristic of Aeromonas is consistent with our finding. We observed the same morphology to what was described of Aeromonas. The colonies were smooth, with a glistening surface. As the for the bacteria cell itself, it is a rod-like shape. Our catalase test and oxidase test shows that Aeromonas can produce enzyme catalase and possesses cytochrome c oxidase. Aeromonas is a Gram-Negative bacterium.  It is consistent with the other test we have done, for example, growth on the Vancomycin, growth on the PEA, and the string test. With the similarities in the physiology and morphology, we can conclude that the DNA sequence was accurate, and the unknown bacteria is Aeromonas.

 The tests that were not consistent with our finding is the EMB-lactase test. There was no growth on the EMB-lactose medium. A reason why could have occurred is that while making our live culture, we did not put enough of the cells into the solution. Another inconsistent is the MSA test. There was no growth on our MSA medium which indicates that the bacteria are non-salt tolerance. Aeromonas is a salt-tolerance bacterium but only to about 6%. MSA contains about 7.5% – 10% concentration of salt. This could have been a reason why we had no growth in our MSA medium.

 Overall, most of our results were consistent with the sequencing and the BLAST results. We conclude that the unknown bacteria was Aeromonas, a gastrointestinal pathogen that can 

cause food poisoning and diarrhea.

References

“BLAST: Basic Local Alignment Search Tool.” National Center for Biotechnology Information, U.S. National Library of Medicine, https://blast.ncbi.nlm.nih.gov/Blast.cgi.

“Chromas: Technelysium Pty Ltd.” Chromas | Technelysium Pty Ltd, https://technelysium.com.au/wp/chromas/.

Holbrook, Mark A., Brenda G. Leicht. Diversity of Form and Function Biology 1412 Lab Manual. Eight edition, Department Of Biology at the University of Iowa, 2019

Igbinosa, Isoken H, et al. “Emerging Aeromonas Species Infections and Their Significance in Public Health.” TheScientificWorldJournal, The Scientific World Journal, 2012, https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3373137/.

Sadava, Hillis, Heller, and Berenbaum. Life: The Science of Biology. Tenth edition, Sinauer Associates, 2014.

 

Processes of Gene Regulation in Bacteria and Archaea

Introduction
The daily life of a microscopic organism consists of adapting to a changing environment and altering its gene expression to regulate these changes in its cells. Bacteria and archaea are prokaryotic cells that sense and respond to changes in their surroundings. Gene regulation helps microorganisms compete in unfavourable, competitive environments by regulating the function of macromolecules such as protein to maximize their resources. Gene expression in bacteria is the response to the available nutrient sources, controlling the amount and type of protein or enzyme. The process of gene expression regulates protein synthesis through transcription by controlling the amount of mRNA available, or through translation which decides whether or not to translate mRNA. Enzyme synthesis and enzyme activity allow a cell to control its metabolism through regulatory processes. Chemotaxis is a process that allows motile bacteria to move towards attractants such as nutrients and away from repellents such as toxins (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, pp. 209 – 210). This paper explores methods of gene regulation in prokaryotic cells such as the lac operon, global control mechanisms of catabolite repression, omp system, alternative sigma factors, Chemotaxis systems, and quorum sensing. It also discusses the interaction of protein-nucleic acid, endospore formation, flagellar rotation and adaptation, and virulence factors in relation to these systems.
Protein-Nucleic Acid Interaction
Regulatory proteins are binding proteins responsible for regulating gene expressions and allow transcription to be turned on and off. A specific promoter on DNA must be recognized to allow a gene to be transcribed. Transcription is a microbial regulation mechanism used by prokaryotic cells. The location protein-nucleic acid interaction occurs depends if the binding protein has a specific binding site or not on the nucleic acid. The protein-nucleic acid interactions are essential to the gene regulation processes of replication, transcription, and translation. A DNA-binding protein; helix-turn-helix is formed of a polypeptide chain with two segments. A short sequence is connected to an α-helix secondary structure, which creates the turn in the structure. This helix-turn-helix structure used by DNA-binding proteins in Bacteria such as Escherichia coli (E. coli ) have lac and trp repressor proteins (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, pp. 210 – 212).
Lac Operon
Negative control prevents transcription by repressing mRNA synthesis, which is the first step in the flow of biological information. A gene that is transcribed more often will have a larger amount of protein available in a cell, resulting in more RNA of that gene available for translation. If products such as amino acid arginine, used by E. coli are present in sufficient amounts in a medium, enzymes that catalyze the synthesis are not produced, they are repressed. Enzyme repression affects biosynthetic (anabolic) enzymes, the opposite of repression is induction which affects degradative (catabolic) enzymes. Induction occurs when a substrate is present and produces enzymes. The sugar lactose is used by E. coli as a source of carbon and energy. β-galactosidase is an enzyme that cleaves lactose into glucose and galactose and is necessary for E. coli to form on lactose. Synthesis of β-galactosidase is not induced until lactose is added and is encoded by the lac operon to allow bacteria to use lactose as a source of energy. The lactose operon, known as the lac operon, is used to ferment lactose. This is a control mechanism used to synthesize enzymes only when necessary (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 212).

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An inducer and corepressor are substances that induce and repress enzyme synthesis, respectively. These small molecules are known as effectors and are not substrates or end products of the synthesis. These substances affect transcription by indirectly binding to DNA-binding proteins. Arginine genes induce transcription from the arginine operon when cells need arginine and repress when the enzyme is abundant. Arginine binds to the corepressor in a cell and activates the protein, where the repressor protein can bind to the operator; a DNA region close to the promoter of the gene. This is a region of consecutive genes called the operon, controlled by one operator. The promoter initiates synthesis of mRNA, upstream from the operator. Transcription can be blocked if the repressor binds to the operator and the encoded polypeptides are repressed and not synthesized. When an inducer is absent, a repressor is active and can control induction, blocking transcription. The repressor protein becomes inactivated when an inducer is added and the effectors combine, overcoming inhibition and allowing transcription to continue (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 213).
Positive control activates transcription. The activator is a regulatory protein that binds RNA polymerase to DNA. In E. coli, maltose is synthesised by enzymes after maltose is added to the medium. The sequence of these enzymes is the same as β-galactosidase as mentioned above, using maltose instead of lactose to induce gene expression. Except, the synthesis of maltose-degrading enzymes is under positive control, instead of negative control with the lac operon, requiring an activator protein to bind to DNA. The maltose activator protein must first bind the maltose inducer before it can bind to DNA. RNA polymerase can then start transcription. Operons of Bacteria and Archaea have many control features; therefore, a network of interactions is vital for regulation (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, pp. 213-214).
Global Control Mechanisms of Catabolite Repression
Global control systems are regulatory mechanisms that regulate gene transcription as a response to a signal in an environment. The lac operon responds to global controls and their own negative and positive controls. Multiple usable sources of carbon may be present for bacteria, E. coli can use various sugars. Catabolite repression is a global control mechanism that chooses the best source of carbon to be used if more than one is present. When glucose is available, it is used first because E.coli forms quicker on glucose compared to other sources of carbon. Catabolite repression uses the best source of carbon and energy first for an organism and is known as the glucose effect. The synthesis of enzymes used in breaking down carbon sources other than glucose are repressed when growing E. coli in glucose-containing medium. Induction is therefore repressed by the presence of the best source of carbon. Global control of the catabolite repression indicates the use of the best source of energy by preventing other catabolic operons, such as lactose and maltose from being expressed while glucose is available. Genes of flagella synthesis are also controlled by catabolite repression due to bacteria having a sufficient source of carbon and not having to swim to find other nutrients. Diauxic growth is a potential consequence of catabolite repression. When two usable sources of energy are present, the better source; glucose will be consumed first, the cell will stop growing when the resource is depleted. After a lag period, the cells use the second energy source; lactose, cells grow faster with glucose than lactose. The synthesis of proteins used by the lac operon rely on catabolite repression and is not expressed in the presence of glucose, lactose is not used. β-galactosidase is necessary for including cells to use lactose. Abolishment of catabolite repression occurs when the glucose is depleted and lac operon is expressed, cells can then use lactose to grow (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 215).
Catabolite repression depends on a regulatory protein that acts as a positive control, called cyclic AMP receptor protein (CRP). RNA polymerases bind to a promoter if CRP binds to DNA and gene encoding catabolite repression enzymes are expressed. DNA must first bind to cyclic adenosine monophosphate (cyclic AMP, also known as cAMP)before it can bind to CRP. Cyclic AMP is a regulatory nucleotide and is synthesized by adenylate cyclase from ATP while its synthesis can be disrupted by glucose, potentially stimulating the transport of cyclic AMP outside the cell wall. The level of cyclic AMP is lowered when glucose is added to the cell. Polymerase can not bind to promoters of catabolite repression when levels of cyclic AMP are low due to CRP not being able to bind DNA. A low concentration of cyclic AMP influences catabolite repression, which is impacted due to glucose being a better source of energy. Transcription of the lac operon relies on the absence of glucose and a high concentration of cyclic AMP. This allows CRP to bind to the CRP-binding site, and an inducer such as lactose must prevent the lactose repressor binding to the operator, blocking transcription (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, pp. 216 – 217).
Omp System
Two-component regulatory systems contain a sensor kinase protein and a response regulator protein located in the cytoplasmic membrane and cytoplasm, respectfully. Autophosphorylation is a process used by sensor kinases; histidine kinases enzymes, to phosphorylate themselves when an environmental signal is detected. Kinase uses phosphate from ATP to phosphorylate compounds and transfers the protein from a sensor into the cell to the response regulator; another protein. Transcription is regulated by a DNA-binding protein as the phosphorylated response regulation functions either act as an inducer by allowing transcription or acts as a repressor, binding DNA and blocking transcription. If repressed, transcription can occur when the response regulator is released and dephosphorylated. A feedback loop is required for a regulatory system to operate correctly. This system uses a phosphatase enzyme to remove phosphate from the response regulator and terminate the response, resetting the cycle of the system (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 219).
There are close to 50 two-component systems in E. coli. The levels of OmpC and OmpF in the outer membrane of E. coli are controlled by the osmolarity of the environment. OmpC and OmpF are porins, proteins that are part of the Omp system, allowing metabolites to pass through the outer membrane of Gram-negative bacteria. Synthesis of these proteins depends on osmotic pressure. OmpC is a porin that has a small pore; synthesis is larger in quantities if pressure is high, and OmpF is a porin that has a large pore; synthesis increases when pressure is low. Located in the cytoplasmic membrane is a sensor histidine kinase called EnvZ, it detects osmotic pressure changes and moves phosphate to OmpR, the system’s response regulator. OmpR-P; phosphorylated OmpR, activates transcription of ompF gene in low osmotic pressure conditions. Omp-R activates transcription of OmpC in high-pressure osmotic conditions and represses transcription of the ompF gene. Regulatory RNA is a control mechanism that regulates the expression of ompF (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 219).
Alternate Sigma Factors
Complex transduction systems exist that contain multiple regulatory elements. Nitrogen assimilation in bacteria is regulated by the Ntr regulatory system. Nitrogen regulator I (NRI) is the response regulator that activates transcription from promoters using the alternative sigma factor s54 (RpoN) to recognize RNA polymerase. In the Ntr system, a sensor kinase; nitrogen regulator II (NRII), is a protein regulated by a second protein called PII, who in turn, is regulated by the contribution of uridine monophosphate (UMP) groups. NRII acts as a kinase and a phosphatase, and the activity of kinase is promoted by the PII-UMP complex during periods of nitrogen starvation, where UMP connects to PII, resulting in phosphorylation of NRII. Phosphatase activity of NRII is promoted by the removal of UMP from PII (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 219 – 220).
Another two-component regulatory system with more than one regulator is the Nar regulatory system. During anaerobic respiration, the use of nitrate and/or nitrite as alternative electron acceptors are controlled by the Nar system. This system contains two different response regulators and sensor kinases, while fumarate nitrite regulator (FNR), controls all genes regulated by the Nar System. Complex chains of regulating systems such as two-component systems for bacteria to phosphorylate histidine residues are common for systems related to cellular metabolism (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 220).
Stringent response is a regulatory mechanism bacterium use to survive changes in their environment such as stresses, antibiotics, and nutrient deprivation. When stringent responses are triggered, stress survival pathways are activated (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 224). Alternative sigma factor RpoS (σs or σ38) is a general stress response that controls these pathways of Gram-negative bacteria cells, who have stress responses and the stringent responses. Gram-positive cells face environmental stresses by undergoing sporulation. These processes allow cells to adapt to environmental stressors and harsh conditions. RpoS regulates more than 400 genes and is the master controller of gene regulation in bacteria. RpoS regulon sense changes in the environment such as nutrients, biofilm formation and stresses and transfer signals to other regulators such as the heat shock response. The dinB gene of E. coli encodes DNA polymerase IV and is recognized by RpoS (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 226 & 228).
RpoH (σ32) is the alternative sigma factor that controls the expression of proteins in the heat shock response of E. coli. Degradation of RpoH synthesis often occurs within a couple of minutes. When cells are affected by heat shock, degradation is inhibited, and transcription of operons increases as more promoters are recognized. Free DnaK inactivates RpoH, influencing degradation. When proteins unfold due to heat stress, DnaK cannot degrade RpoH due to binding with unfolded proteins. The level of free DnaK is affected by the number of denatured proteins. With less free DnaK, the level of RpoH is higher which causes expression of the heat shock genes. DnaK inactivates RpoH when environmental stressors such as temperature disappear and the reduction of heat shock protein synthesis occurs (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 227).
Endospore Formation
Endospore formation is used by microorganisms as a response to harsh environmental conditions, forming spores from vegetative cells. Spores germinate when the conditions improve, and the organism’s cells return to a state of vegetation. Bacillus spp. are Gram-positive bacteria that form endospores within the mother cell and cells divide asymmetrically before endospore formation. Endospores are released once formed and the mother cells burst. Unfavourable environmental conditions trigger endospore formation of Bacillus subtilis (B. subtilis) which use five sensor kinases to monitor their environment. Sporulation factors are the successive phosphorylation of multiple proteins resulting from responding to cumulative adverse conditions. Spo0A is a sporulation factor that controls the expression of genes specific for sporulation. Sporulation occurs when Spo0A is highly phosphorylated. The σF sigma factor is an important part of the sporulation process. This product requires SpoIIE to remove phosphate from SpoIIAA, triggering the removal of SpoIIAB, the anti-sigma factor (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 248).
Endospore development of B. subtilis are controlled by four σfactors. σF and σG activate genes required for developing forespores, while σE and σK activate genes required for the mother cell that surrounds the endospore. σF is bound by an anti-sigma factor, therefore, inactive in the forespore, and becomes activated by the Spo0A transmitting a spore signal. Transcription is promoted when σF binds to RNA polymerase and produces the gene encoding σG sigma factor. σE is important for transcribing genes for σK and requires to be activated by genes for proteins entering the mother cell, required for the sporulation process. This process also requires sigma factors σG and σK to later transcribe genes. After a long process, spore coats and structures are formed, and the endospore can be released. Sporulation in Bacillus is often triggered by the limitation of nutrients in the environment (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 248).
Chemotaxis Systems 
Chemotaxis regulation is the movement of Bacteria and Archaea cells away from repellants, towards attractants as they respond to changes in the environment such as toxin accumulation and limited nutrients. Prokaryotic cells sense and respond to temporal gradient changes in the concentration of a chemical over time, they are not large enough to sense the absolute concentration of a chemical. Bacteria have a unique, modified two-component system that regulates the direction of flagellar rotation by sensing temporal changes within repellants or attractants. Instead of encoding flagellar proteins by controlling transcription, this system regulates the activity of the flagellum protein complex. Sensory proteins called methyl-accepting chemotaxis proteins (MCPs) interact with cytoplasmic sensor kinases in the cytoplasmic membrane for monitoring concentrations of substances. MCPs sense attractants or repellants which drive the mechanism of chemotaxis through signals from proteins (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, pp. 220 – 221).
The Tar MCP transmembrane production of E. coli sense cobalt and nickel as repellents and maltose and aspartate as attractants. Flagellar rotation is influenced through interactions between MCPs binding attractants or repellents to periplasmic binding proteins which interact with cytoplasmic proteins. Chemoreceptors are made of thousands of clustered MCPs that contact CheA and CheW cytoplasmic proteins. In chemotaxis, the sensor kinase is CheA, and CheW helps with autophosphorylation of CheA to CheA-P after MCP binds a chemical. The concentration of an attractant affects the rate of autophosphorylation; as a concentration increases, autophosphorylation decreases, and with an increase in concentration of repellent or decrease in attractant, increases autophosphorylation. Flagellar rotation is controlled by a response regulator that forms when Phosphate is passed from CheA-P to CheY creating CheY-P. Phosphate can be transferred from CheA-P to CheB, which is important for adaptation (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 221). 
Flagellar Rotation and Adaptation
CheY controls the direction of flagellar rotation and is a central protein of the regulatory system. After phosphorylation occurs, CheY induces clockwise rotation of the flagellum causing the cell to tumble. If CheY is phosphorylated (CheY-P), the motor of the flagellum can not bind and turns counter-clockwise, sending the cell to run; swimming smoothly. Runs occur when CheZ dephosphorylates CheY preventing cells from tumbling. Tumbling relates to an increase in the level of Chey-P, which depends on an increase in concentration of repellent or decrease of attractant. Runs are promoted by moving towards an attractant, tumbling is then suppressed due to the lower level of CheY-P. After responding to stimulus, an organism resets the sensory system through a feedback loop relying on CheB, this is known as adaptation of the chemotaxis system as the organisms must wait for another signal (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 221 – 222).
MCPs are sensitive towards repellents and when methylated, they do not respond to attractants. MCPs respond to attractants and not to repellents when unmethylated. Methylation and demethylation from CheR and CheB-P cause variation in levels of methylation, allowing adaptation to sensory signals. Autophosphorylation rate of CheA is low in relation to a high concentration of attractant. This causes cells to swim smoothly due to unphosphorylated CheY and CheB. At this time, methylation of MCPs increases as they can not be demethylated by CheB-P, as CheB-P is not around. When fully methylated, MCPs do not respond to an attractant causing the cell to tumble due to the high, constant level of attractant. When CheB is phosphorylated, CheB-P can demethylate the MCPs resetting the receptors, allowing them to respond to changes in levels of attractant. When fully methylated, MCPs signal for a cell to begin tumbling as they respond to an increase in gradient of a repellent. MCPs slowly demethylate as cells tumble in random directions. Chemotaxis successfully monitors small changes of concentrations over time in repellents and attractants through this mechanism for adaptation (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 221 – 222).
Quorum sensing
Quorum means sufficient numbers. Quorum-sensing systems in bacteria respond to other cells of the same species around the cell, and the density of cells can control the regulatory pathway. This is known as quorum sensing. Some species of Archaea also contain quorum-sensing systems. Mechanisms of quorum sensing assess density of populations, bacteria use this to make sure the proper cell density is present before operating, correctly. Individual cells of toxin secreting pathogenic bacterium would waste resources as it would not have an effect on a host. Although, coordinated expression toxin-secreting bacteria cell may cause illness, thriving from collecting resources from the host, benefiting the pathogen. An autoinducer is a signal molecule synthesized through quorum sensing used by both Gram-positive and Gram-negative bacteria but is more common in the latter. The molecule diffuses in either direction across the cell envelope, causing the autoinducer to increase in concentration within the cell if multiple cells making the same autoinducer are around. The autoinducer triggers gene transcription binding to a transcriptional activator protein or sensor kinase of a two-component system. Gram-negative bacteria contain different lengths of acyl groups containing carbonyl and alkyl called Acyl homoserine lactones (AHLs). Autoinducer 2 (AI-2) containing cyclic furan derivative is made by some Gram-negative bacteria and is used between bacteria species as a common autoinducer. On the other hand, Gram-positive bacteria use short peptide autoinducers (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 222).
Bioluminescent light emissions in bacteria are regulated by a quorum sensing mechanism. Aliivibrio fischeri is a marine bacterial species that emit light generated by an enzyme called luciferase. Proteins needed for bioluminescence are encoded by lux operons controlled by activator protein LuxR. Bioluminescence is induced by an increase in concentration of N-3-oxohexanoyl homoserine lactone, a specific AHL of A. fischeri. The luxI gene synthesises AHL through the encoded enzyme (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 222).
Virulence Factors
Quorum sensing control genes of pathogenic E. coli 0157:H7, a Shiga toxin-producing foodborne pathogen induces virulence genes by making AI-3. While the population of E. coli cells in the intestine increases; producing AI-3, the stress hormones norepinephrine and epinephrine are made by host cells in the intestine. In the cytoplasmic membrane of E. coli, the signal molecules become bound to sensor kinases. This activates two transcriptional activator proteins which activate transcription of genes that secrete toxins, encode proteins causing lesions on intestinal mucosa of a host, encode the function of motility, and phosphorylation. This system regulates gene expression by sensing chemical signals of both bacterial and eukaryotic cells (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 223).
Autoinducing peptide (AIP) is a small peptide used by the quorum-sensing system to control virulence factors that encode genes and impact the immune system of a host organism. Staphylococcus aureus is a pathogenic bacterium that uses extracellular peptides to affect cells of an organism. The argD gene is the autoinducer that encodes AIP, and after its, synthesis the peptide is trimmed into the active form of AIP by the membrane-bound protein ArgB and the peptide used by S. aureus is excreted outside the cell. Concentration of AIP increases along with cell density of S. aureus. Autophosphorylation occurs from the binding of AIP to the membrane-bound sensor kinase ArgC, producing ArgC-P and transfers its phosphate to a transcriptional activator called ArgA, producing ArgA-P. Production of virulence proteins is controlled by an RNA molecule transcribed by argABCD genes. Transcription of these genes is influenced by an increase of ArgA-P, encoding the signal transduction systems. Eukaryotic organisms have been known to disrupt quorum sensing in bacteria by producing molecules that mimic AHLs or AI-2, interfering with the behaviour of bacteria. These quorum-sensing disruptors can potentially be used to impede virulence gene expression and disperse bacterial biofilms (Madigan, Bender, Buckley, Sattley, & Stahl, 2018, p. 223).
Conclusion
Regulatory mechanisms of gene expression are used by prokaryotic cells to sense signals in their environment and bind to DNA. Prokaryotic cells respond to environmental sensors and stressors by turning genes off and on. Endospore formation is used as in Gram-positive bacteria to enter a state of dormancy when unfavourable conditions arise. It is important to understand how these systems work to ensure the correct gene is present and it can be turned on in the available conditions. Understanding how these systems work and how they work with microorganisms in their natural environment is vital for bioremediation and designing applications for industrial use (Noble, 2021).
References
Madigan, M. T., Bender, K. S., Buckley, D. H., Sattley, W. M., & Stahl, D. A. (2018). Brock biology of microorganisms (15th ed.). New York: Pearson.
Noble, N. M. (2021). Lecture 1 – How do cells sense the environment.doc. Course Notes and Lecture. ENSC317. Victoria BC: Royal Roads University. https://moodle.royalroads.ca/moodle/mod/forum/discuss.php?
 

Bacteria, Viruses, and Parasites in Philippines Water

Resorts in the province of Cavite are recognized for its reputable recreational waters. These resorts are intended to provide prospective customers with an atmosphere of amusement, entertainment and relaxation. The most common types are beach resorts, swimming pool, and even lakes and rivers which are designed to accommodate individuals, group of peers and family members (Bago and Linantud 2004). Also recreational waters offer activities that are beneficial and substantial to overall health.

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Recreational waters can be contaminated and polluted by bacteria, viruses and protozoan parasites (Bitton 1999), although the recreational water is normally treated physically and chemically using filtration and chlorination to prevent growth and infection of some bacteria (Montano and Abear 2000). However there are was an increasing number of cases of acute gastroenteritis during this summer and one group of microbes leading to their disease are enteric bacteria.
Bacteria such as Escherichia coli and Pseudomonass aeruginosa that are resistant and tolerant to chlorine and were known to cause human misery (Mann 2005). Enteric Bacteria are said to be notorious and dangerous because they cause recreational water illness like acute gastroenteritis, cholera, pneumonia, typoid fever, diarrhea, urinary infection, pneumonia, dermatitis, salmonellosis and otitis external these disease leads to outbreaks (Yoder 2008). Escherichia coli and Pseudomonas aeruginosa are among those bacteria that can thrive in recreational waters and transmitted from swallowing and inhaling contaminated water before the microorganisms can be destroyed by pool water disinfectant (Barwicks et al. 1999). Also their resiliency to grow in some disinfectants like chlorine because they are capsulated bacteria and they possess a versatile metabolic activity, which makes gives them the resistance to a variety of physical conditions (Cappuccino 2005). The presence of these organisms indicates contamination by pathogenic microorganism. Most waterborne diseases are related to pollution of water resources sources and thus pose an unacceptable health risk for swimmers (Schets et al. 2010). Therefore the need to examine water samples in a microbiological water aspect is essential to ensure safety to swimmers.
This study is conducted to support if the selected resorts in Cavite whether chlorinated and non-chlorinate is contaminated with pathogenic microorganism that can lead to potential waterborne diseases.
1.2 Conceptual Framework
The water districts ensure the potability of drinking water by increasing the concentration of Chlorine (Yoder 2008). This potable drinking water were also used in resorts. Monthly sampling of water samples in pools render negative in enteric bacteria. However there is an increasing cases of gastroenteritis for the past year.
The paradigm of the present study is as follows :
Microbial Status in Chlorinated and non- chlorinated waters
from selected resorts
Water samples from resorts (pools, streams, rivers and beaches)
ed
The objective of this study is to gather different water samples obtained from selected resorts and determine the microbial status regardless of its chlorine concentration levels.
1.3 Statement of the Problem
This study will aim to determine the microbial status in selected resorts in Cavite.
To determine specifically the following objectives:
1. What is the microbial status jpresent in chlorinated and non -chlorinated water in selected resorts in Cavite?
2. Is there a significant difference in the total count of enteric bacteria and total bacteria among chlorinated and non-chlorinated waters in selected resorts in Cavite?
3. Is there a correlation between chlorine concentration in the resorts and water samples in lakes, rivers and beaches?
1.4 Scope and Delimitation
The study will determine microbial status in selected resorts in Cavite, Province. Water samples will be taken from these selected resorts and chlorine content concentration will be determined using chlorine test kits. It is not the aim of the study to apply antibacterial agent in enteric bacteria recovered from water analysis.
1.5 Significance of the Study
This study is designed to find out the microbial status on the selected resorts in Cavite, Philippines. This research hopes to benefit the following concerned population:
Resort clientele to be more concerned to the areas that they went to and be prepared since traditional vaccines are not reliable in killing these bacteria because they are risky and are only effective after several years.
Resort Administrators and Maintenance personnel for the enhancement of facilities of the swimming pool to promote the preventive measures against proliferation of microorganism which are recognized to be health risk problems and compliance to the water quality standards.
Academe who may use this as preliminary information for their future research endeavors and information in survival of bacteria in different environmental conditions.
1.6 Definition of Terms
Microbial status – this refers to the bacteria present in chlorinated and non-chlorinated waters from selected resorts.
Prevalence – the number of samples that rendered positive in culture method over the total number of samples.
Enteric Bacteria – these are large group of gram-negative bacteria that are known to produce disease in the alimentary tract. Enteric bacteria that survived in chlorinated waters of resorts.
Resorts – it is considered to be swimming pools, streams, river, lakes and beaches with chlorine.
Microbial Density- The population or the measurement of the growth of the bacteria.
Microbial Plating- This refers technique used to isolate a pure strain from a single species of microorganism plating method that will be performed in laboratory.
Total Bacteria – bacteria other than enteric bacteria.
API Kit- This refers to the biochemical test that will determine the isolated bacteria from water samples.
Chlorination- this is a water purification method to make water safe to humans and a disinfecting agent that prevents the spread the spread of waterborne diseases
Chlorine test kit- This refers to the chemical test that will determine the chlorine concentration level of water.
Chromogenic Media – This refers to the culturing media that will determine the present bacteria in water.
CHAPTER 2
REVIEW LITERATURES
2.1 Conceptual Literatures
Recreational water
Recreational waters can be classified as fresh water swimming pools, whirlpools and naturally occurring fresh marine surface waters. Infectious disease which can be transmitted by recreational water includes skin, eye and ear infections and gastroenteritis. Consequently the level of microorganism in recreational water are important for indexing their health hazard associated with swimming and since the recreation classification includes bathing, swimming etc. any organism transmitted to humans can be regulated. The best indicators in the assessment of the safety of swimming pool water is to become aware of the types of hazard (microbiological, chemical and physical) that can impact a bathing area. Some researchers emphasize that the microbiological quality of swimming pools are best measure by identifying the bacteria present in that recreational water such as fecal coliform and enterococci, while others consider that the disease and symptoms it brought to the bathers rather than fecal contamination (Martin et al. 1995).
(Montano and Abear 2000) cited that the bacteria suggested as indicators of recreational water quality include a wide variety of pathogenic bacteria and non-pathogenic microorganism such as coliform groups, species of Pseudomonas, Streptococcus, Staphylococcus and in rare case Legionella. The presence of single coliform organism is not a ground for condemning water as a unit for human consumption. It is the relative abundance of these organisms, which is important.
According to Papadopoulou et al. (2007) cited that non-fecal human shedding (e.g. from vomit, mucus, saliva or skin) in the swimming pool is also a potential source of pathogenic organism. Bathers who are already infected can directly contaminate pool waters with pathogen which may affect other bathers, who come in contact with the contaminated water. ‘Opportunistic pathogens’ (mainly bacteria) can also be shed from user and transmitted via contaminated water. Also certain free living aquatic bacteria and amoebae can possibly grow not just in pool waters but also with pool components or facilities or on other wet surfaces within the facility which may cause infections or disease. Therefore swimming pools are often associated with outbreaks or incidents of waterborne infection.
Murdoch(1975) as cited by Amador and Amante (2001) mentioned that disease contracted from water kill some 25million people, most of them children each year, while many millions more are debilitated by waterborne diseases. Fecal contamination of water can introduce a variety pathogens into water waste, including bacteria, viruses, protozoans and parasitic worms. Waterborne related diseases have been recognized by Classes. Class 1, refers to the true waterborne disease contracted by drinking water. Class 2 are diseases associated with lack of personal hygiene which can be reduced by providing adequate amount of water for bathing and washing. To control such diseases, people should be provided with sufficient water of reasonable quality; achieving a high bacteriological quality is a secondary consideration.
Enteric bacteria
A large, heterogenous group in the family Enterobacteriaceae, include several closely related genera of short and spore forming, gram-negative rods, facultative anaerobic, that inhabit or produce disease in the alimentary tract of warm-blooded animal. This family are notorious as causes of urinary tract infection and are recovered from a variety of clinical specimens taken from diseased foci other than in the gastrointestinal tract. The enterobacteria are probably responsible for more human misery than any other group.(Smith 2008)
Escherichia coli
It is a gram negative rod shaped bacterium. It was originally known as bacterium coli. It is widely distributed in the intestine of humans and warm- blooded animals and is the predominant facultative anaerobe in the bowel part of the essential intestinal flora that maintains the physiology of the healthy host. The presence of E.coli is associated with bather-associated illness, but its absence cannot be equated with the lack of risk of illness (Guidelines for Canadian Recreational Water Quality available at http://www.ecy.wa.gov1992).
Pathogenicity performs coliform bacilli usually do no penetrate intestinal wall to produce disease unless (1) the intestinal wall becomes diseased, (2) resistance of the host is lowered, or (3) virulence of the organism is greatly increased. Under one of these conditions of coliforms may pass to abdominal cavity or enter into the bloodstream. Once outside the intestinal canal and in the tissues of the body their virulence is remarkably enhanced. Among the diseases that they cause are pyelonephritis, cystitis, cholecystitis, abscesses, peritonitis, and meningitis. They may play a part in the formation of gallstones and are found in the cores of such stones. In peritonitis complicating intestinal perforation the coliform group is joined by such organisms as streptococci and staphylococci. From any focus of inflammation coliform organism may enter the bloodstream to produce a septicaemia. (Smith 2008)
Shigella
Dysentery caused by the Shiga bacillus (Shigella dysenteriae) is much more severe than that from the other organisms, since this bacillus produces a powerful exotoxin- like substance in addition to an endotoxin. The exotoxin- like substance seems to be liberated by bacterial disintegration, and as a neurotoxin, It acts on the nervous system to paralyze the host. The endotoxin irritates the intestinal canal.
The dysentery bacilli are gram negative, nonsporebearing rods that grow on all ordinary media at temperatures from 10° to 42° C. but best at 37° C they are aerobic and facultative anaerobic. Unlike most other members most other member of the enteric group, they are non-motile.
In terms of pathogenicity dysentery is a human disease and natural infections of the lower animals do not occur. The incubation period is 1 to 7 days. Epidemic dysentery is primarily an intestinal infection. Unlike typhoid bacilli, the organisms do no invade the bloodstream and are seldom if ever found in the internal organs or excreted in the urine. They are excreted in the feces. Compared to that for other enteric pathogen, the number of ingested shigellas for infection is small, only 10 to 100. (Smith 2008)
Salmonella
Among the large number of pathogenic microorganisms causing foodborne disease, Salmonella plays an important role. An analysis of Salmonella surveillance data from the World Health Organization (WHO) showed that the reported number of cases increased in 22 out of 49 countries examined. Although the reason for the global increase is not yet clear, investigations in individual countries suggest that it is related to consumption of eggs and poultry that harbour the organism. Besides control measures there is a need for rapid and sensitive methods for the detection of Salmonella (Beumer et. al, 1991). Salmonella is a ubiquitous enteric pathogen with a worldwide distribution that comprises large number of serovars characterized by different host specificity and distribution. This microorganism is one of the leading causes of intestinal illness through the world as well as the etiological agent of more severe systemic diseases such as typhoid and paratyphoid fever.
Zoonotic salmonellae are commonly described as foodborne pathogens however; drinking water as well as natural waters is known to be an important source for the transmission of these enteric microorganisms. Salmonella, just like other enteric bacteria, is spread by the fecal-oral route of contamination. This microorganism can enter the aquatic environment directly with feces of infected humans or animals or indirectly, e.g., via sewage discharge or agricultural land run off.
Overall Salmonella spp. and subspecies can be found in a large variety of vertebrates. Beside humans, animal sources of Salmonella include pets, farm animals and wild animals; calves, poultry, pigs, sheep as well as wild bird (pigeon) and reptiles can all be reservoirs of Salmonella. Plants, insects and algae were also found capable of harboring Salmonella and might be implicated in the transmission of this enteric pathogen. Taxonomically the genus Salmonella comprises two species namely S. bongori and S. enterica. The species S. enterica is further differentiated in to six subspecies (enterica, salamae, arizonae, diarizonae, indica and houtenae) among which the S. enterica subspecies enterica is mainly associated to human and other warm blooded vertebrates. Enteric fevers, typhoid and paratyphoid fever are severe, contagious systemic diseases caused by the infection of the serovars typhi and Paratyphi. Differently from other Salmonella serovars, typhi and Paratyphi are host adapted and can only infect humans; stools of infected persons are therefore the original source of contaminations for these pathogens.
Water contaminated with feces of human cases and carriers is one of the main vehicles of typhoid fever infections. Literature data related to water-borne salmonellae in developing countries relate mostly the typhoid Salmonella serovars. In the less industrialized area of the world, in particular in the Indian subcontinent and South East Asia, typhoid and paratyphoid fevers occur both in epidemic and endemic form, and remain a major public health problem. The burden of typhoid fever worldwide is further compounded by the spread of multiple drug resistant S. typhi.
Most of the recent publications on typhoid and paratyphoid fever water-borne infections in developing countries are from the Asian continent. Differently from typhoidal Salmonella strains, non-typhoidal salmonellae, the ubiquitous subtypes found in a number of animal species, are more frequently associated to foodborne than to water-borne transmission. These zoonotic Salmonella serovars tend to cause acute but usually self-limiting gastroenteritis (Levantesi et al, 2011).
According to (Smith 2008)The pathogenicity of salmonella is called salmonellosis, the major site of which the lining of the intestinal tract. Because of their toxic properties every known strain of salmonella can cause anyone three types of salmonellosis: (1) acute gastroenteritis of the food type infection.(2) septicemia or acute sepsis with localized complications similar to pyogenic infections, and (3) enteric fever such as typhoid or paratyphoid fevers.
Salmonella typhi
A short motile nonencapsulated bacillus, S.typhi grows luxuriantly on all ordinary media. It grows best under aerobic conditions bit may grow anaerobically. The temperature range growth is from 4° to 40°C., the optimum, 37°C. typhoid bacilli can survive outside the body, living about 1 week in sewage contaminated water and not only living but multiplying in milk. They may be viable in fecal matter for 1 or 2 months. They are pathogenic because of their endotoxins.
Their pathogenicity causes typhoid fever is an acute infectious disease with continuous fever, skin eruptions, bowel disturbances, and profound toxemia. Except in the first few days, leukopenia is always present in uncomplicated cases, probably because typhoid bacilli depress the bone marrow, where normal production of white blood cells occurs. Leukocytosis in the course of the disease signals complication. (Smith 2008)
2.2 Related Studies
According to Brown (2009), gram-negative intestinal pathogens have a diverse population of bacteria of which two of the enteric intestinal pathogens that are of prime medical concern are the salmonella and shigella. The salmonella and shigella are both pathogenic bacteria that cause typhoid fever and human dysentery, respectively. Since the gram-negative intestinal pathogens has a such diverse population it has many genera of species like the Escherichia, Proteus, Enterobacter, Pseudomonas, and Clostridium that exists on large numbers, hence it is necessary to use media that are differential and selective to favor the growth of the pathogens since all of the species can be divided into lactose fermenting and non-lactose fermenting bacteria.
Hiriart et al. (2001) worked on the Helicobacter pylori and Other Enteric Bacteria in Freshwater Environments in Mexico City. They observed that all samples analyzed showed the presence of enteric bacteria with or without the presence of H. pylori, indicating that water from these sources is a potential health risk for gastrointestinal diseases. The major positivity of H. pylori coincides with the major positivity of indicator and other enteric bacteria, which are both associated with contaminated water.
In another study Marion et al. (2010) worked on the association gastrointestinal illness and recreational water exposure at an inland U.S beach. Relationships between water quality indicators and reported adverse health outcomes among users of a beach at an inland U.S lake was observed to be a significant risk factor for GI illness.
.
Papadopoulo et al.(2008) worked on the microbial quality of indoor and outdoor swimming pools in greece. They found out that three indoor swimming pools and two outdoor swimming are present with bacteria, protozoa and fungi Such as Multi-resistant Pseudomonas alcaligenes, Leuconostoc, and staphyloccus aureus( isolated from teaching pool), Staphylococcus werneri. Chryseobacterium indologenes and Ochrobactrum anthropic (isolated from completion pools) Pseudomonas aeruginosa, P. fluorescens, Aeromonas hydrophila, Enterbacter cloacae, Klebsiella pneumonia and S. aureus (isolated from the hydrotherapy pool and A. hydrophilla (isolated from the hotel pool) were related to water outbreaks.
Schets et al. (2010) worked on the exposure assessments for swimmers in bathing waters and swimming pools. they found out that the swallowed volume or water appears different for men, women, and children, but also in fresh water, seawater and swimming pools also the frequency and duration of swimming do also differ for men, women, and children and in different water types, and provide a basis for the identification of high risk population under specific circumstances, e.g. due to their extended water contact and frequent head submersions, children may be more prone to contract otitis external due to Pseudomonas aeruginosa infections.
Certainly a waterborne infection depends on the total bacterial counts, the immune status of the subjects, and polluted waters. The results of the past studies demonstrate the variability of the recreational water quality and the need for continuous monitoring.
Chapter 3
METHODOLOGY
Research Design
This study will use descriptive study design that involves in the identification of enteric bacteria in selected resorts in the Cavite province. There will be 20 sampling sites, 10 from swimming pools, 5 from rivers or lakes and 5 from beaches. In every sampling site there will be a total of 1 sample that will be gathered and it will be replicated into three and a total of 60 sterilized bottles with cover will be used for the 4-month period of experiment that will be done during the summer season and the rainy season.
Research Setting
The entire study will be conducted for 12 weeks. The identification of total bacteria and enteric bacteria will be done in Biology Research Laboratory of DLSU-D.
Research Procedure
Water Sample Collection (MicroMed Environmental, 2010)
Sterilized 300ml wide-mouthed glass will be used in the collection of samples. Water samples will be obtained from recreational waters. The sterile containers will be plunge into the water surface until 1 foot below. Then open the bottle towards the direction of the current to allow the container to fill. Afterwards, it will be immediately sealed tightly and placed on a cooler to maintain the temperature. The samples will be obtained during the months of april and june of 2012. The chlorine concentration will also be measured using Hach Test Kit for chlorine.
Chromogenic Media for Bacteria
Undiluted samples will be used in the determination of total bacteria. Briefly one milliliter of sample will be spread plated onto Plate Count Agar. The plates will be incubated at 37°C for 24 hours. Colonies that will grow will be converted into colony forming units and will be correlated to chlorine concentration and compared to enteric bacteria.
For the detection of enteric bacteria the samples will be enriched in buffered peptone water for 24 hours. After 24 hours the enriched samples will be spread plated onto Salmonella-Shigella Agar and Eosin Methylene Blue Agar. Colonies resembling to enteric bacteria will be purified and confirmed using API 20E kit.
Determination of the Microbial Count (BioMérieux, 2002)
Preparation of incubation box and inoculum will be done for the strip. In the inoculation of the strip, filling both tube and cupule of tests CIT, VP and GEL with bacterial suspension as for the remaining tests fill only the tube and not the cupule. In creating anaerobiosis ADH, LDC, ODC, H2S and URE should be overlay with mineral oil. The incubation box will be incubated for 37°C for 24 hours. Certain color reactions will happen for the indication of positive or negative result.
Data Gathering
Colonies in the EMBA and PCA will be characterized using colonial characterization which includes size, form, margin, elevation, consistency, surface and pigmentation (Tabo, 2005). Biochemical test include ONPG, ADH, LDC, ODC, CIT, H2S, URE, TDA, IND, VP, GEL, GLU, MAN, INO, SOR, RHA, SAC, MEL, AMY, ARA, OX. The chlorine concentration will be measured in 0-600 mg/L.
Statistical Treatment
To determine the correlation between chlorine concentration and total bacteria and enteric bacteria, a simple correlation will be used. All statistical analysis will be conducted in STATA 9.0 with 0.05 as level of significance.
APPENDIX A
GANTT CHART
APPENDIX B
BUDGET PROPOSAL
Item
Volume/Mass
Estimated Price (PhP)
Quantity
Expense (PhP)
EQUIPMENTS AND KITS
 
 
 
 
Biomerieux Inc Biomerieux API 20E KIT 100g Pack of 100 20160

13000.00
1
13000.00
Hach’s Chlorine Test Strips, 0-600mg/L Pack of 2890200

876.31
1
876.31
AGARS
 
 
Salmonella-Shigella Agar
50 g
500.00
1
500.00
Eosin Methylene Blue Agar
50 g
500.00
1
500.00
Plate Count Agar

500.00
1
500.00
TOTAL
15376.31
APPENDIX C
LETTER TO THE HOSPITAL
March 13, 2012
Ms. Teresita E. Guevarra
Medical Records Head
De La Salle University Medical Center
Dear Ms. Guevarra:
Greetings in the name of St. John Baptist De La Salle!
We are writing to ask permission from you in getting information that we will need for our thesis defense on the upcoming December 2012. We are Human Biology major students from De La Salle University-Dasmariñas and we are going to conduct a study regarding the possible prevalence of enteric bacteria in selected resorts in Dasmariñas, Cavite. Regarding this, we would like to request for the following information:
Reported cases of salmonellosis and acute gastroenteritis in this hospital for the last two years (2010 and 2011)
We are hoping for your positive response towards our request. If ever the information we need will not be available today, you may contact us at 09164745448 and 09272546946. Thank you very much for your time.
Sincerely,
Ron Matthew A. Flores
John Paul A. Flores
Noted by:
____________________ _____________________
Mrs. Hazel Ann L. Tabo Dr. Carmelita C. Cervillon
BSD Faculty, DLSU-D (Thesis Adviser) College Dean, DLSU-D
_____________________
Ms Cherry Z. Cuevas, MS
BSD Chair, DLSU-D
APPENDIX D
COLOR REACTION
TESTS
– RESULTS
(negative)
+ RESULTS
(positive)
ONPG
colorless
yellow
ADH
Yellow
red/orange
LDC
Yellow
red/orange
ODC
Yellow
red/orange
CIT
pale green/yellow
blue-green/blue
H2S
colorless/gray
black deposit
URE
Yellow
red/orange
TDA
Yellow
brown-red
IND
Yellow
red (2 min.)
VP
colorless
pink/red (10 min.)
GEL
no diffusion of black
black diffuse
GLU
blue/blue-green
yellow
MAN
blue/blue-green
yellow
INO
blue/blue-green
yellow
SOR
blue/blue-green
yellow
RHA
blue/blue-green
yellow
SAC
blue/blue-green
yellow
MEL
blue/blue-green
yellow
AMY
blue/blue-green
yellow
ARA
blue/blue-green
yellow
OX
colorless/yellow
violet
LITERATURE CITED
Amador RM, Amante PP. Detection and isolation of coliform bacteria in Laguna de Bay Brgy. Landayan San Pedro Laguna; 2001. p.67.
Bago CEM, Linantud JF, Ortiz MP. Stability and Profitability of Resort Business in Dasmarinas, Cavite. 2004. P.1-2-ix-29.
Barwicks RS., Levy DA., Craun GF., Beach MJ., Calderon RL. 2000.
Surveillance for water borne-Disease Outbreaks-united-states ,1997-1998 CDC
Brown, A. E. 2005. Benson’s Microbiological Applications 9th Edition, McGraw Hill, New York.
Beumer, R.R., et al., 1991. Enzyme-linked immunoassays for the detection of Salmonella spp.: a comparison with other methods, Elsevier Science Publisher, B.V. 0168-1605/91
Carteciano JA., 2004. Four Emerging Bacteria: So Tiny, So deadly. National
Research Council of the Philippines.
Hammer Sr. M, Hammer Jr. M. Water and waste water technology. New Jersey; 2004.p.140
Levantesi, C., et al., 2011.Salmonella in surface and drinking water: Occurrence and water-mediated transmission, Food Research International, doi:10.1016/j.foodres.2011.06.037;
Mann, D. Beware of Recreational Water Illnesses, WebMD. [Internet]. 2005 [cited 2011 December 28].
Available from HYPERLINK “http://www.webmd.com/fitness-exercise/features/beware-of-recreational-water-illnesses”
Marion, J., et al., 2010.Association of Gastrointestinal illness and recreational water exposure at inland U.S beach, water research international;
Martin, M., et al., 1995.Assessment of microbiology quality for swimming pools in South America.
MicroMed Environmental, Inc. [Internet]. 2010 [cited 2012 March 25]. Available from HYPERLINK “http://www.igmicromed.com/docs.html”
Montano JM, Abear R. 2000.Detection of Pseudomonas aeruginosa in relation to microbial population of selected swimming pools in dasmarinas cavite. De la Salle University Dasmarinas. p.52.
Schets F., et al., Exposure Assessment of swimmers in bathing water and swimming pools, water research. 2010.
Tabo, Norbel A. 2005. Laboratory Manual in Microbiology, Rex Bookstore Inc, Manila. p. 63-67
Yoder JS., Hlavasa MC., Craun GF., Hill V., Roberts V., Yu PA., Hicks LA., Alexander NT., Calderon RL., Roy SL., and Beach MJ.2008. Surveillance for waterborne disease and outbreaks associated with recreational water use and other aquatic facility- associated health events-united states 2005-2006- CDC.
 

Antibiotic Resistance in Bacteria Essay

Antibiotic resistance is a serious matter which should be addressed seriously.
Every time you take antibiotics you don’t need you increase your chance of contracting an infection that is caused by bacteria that are resistant to antibiotics. And if you get an infection that can’t be treated by antibiotics you run the risk of your infection getting considerably worse and you might need to be treated in hospital. There are many factors affecting as to how antibiotic resistance acquires but one thing is for sure, it must be stopped!
At present antibiotic resistant poses as a massive challenge for modern medicine. There is a wide variety of conditions that antibiotic resistance stands in the way of successful treatment like tuberculosis (TB) and Methicillin-resistant Staphylococcus aureus (MRSA).
As we know TB is a disorder affecting the lungs and also the rest of the body. It is caused by mycobacterium tuberculosis and it is reported that a third of the world’s population has been infected with mycobacterium tuberculosis. New infections occur at a astonishing rate of one per second. The proportion of people who become sick with tuberculosis each year is stable or falling worldwide but, because of population growth, the absolute number of new cases is still increasing. Prevention relies on screening programs and vaccination(http://who.int/mediacentre/factsheets/fs104/en/index.html)

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Antibiotic resistance is a growing concern in multi-drug-resistant TB. In a scientific journal titled ” Tuberculosis resistant to isonazid and rifampin” published in 1993 it was concluded that patients with tuberculosis that is resistant to isonazid and rifampin often didn’t succomb to the best treatment available and that failure to obscure this reistance would end in high mortality rates and a gloomy reality for the public (Goble et al) .
In the case of Staphylococcus aureus where, like tuberculosis, it has grown resistance to it’s most of it’s treatment. Staphylococcus aureus is very difficult to treat if contracted due to its high resistance rate to a group of antibiotics called beta-lactams. This group of antibiotics includes penicillin, cephalosporins, tetracyclines, clindamycin and vancomycin.There are different treatments for different variations of the disease but treatment isnt straight forward due to the high rate of antibiotic resistance.
In a scientific report titled “High prevalence of multidrug-resistant MRSA in a tertiary care hospital of northern India”, where they were testing the resistance percentages of the known antibiotic treatments on a group of 783 patients, they found that nearly all the antibiotics that were tested, there was a high rate of resistance. For instance, from the 783 patients isolated who had staphylococcus aureus, 301 (38.44%) had shown to be methicillin-resistant, of which 217 (72.1%) were found to be multidrug-resistant. Practically all MRSA strains were showing resistance to penicillin, 95.68% showed resistance to cotrimoxazole, 92.36% showed resistance to chloramphenicol, 90.7% showed resistance to norfloxacin, 76.1% showed resistance to tetracycline, and 75.75% had shown resistance to ciprofloxacin. The antibiotic showing the least amount of resistance was vancomycin with 0.33%. (Hare Krishna Tiwari et al).
How bacteria become resistant
The actual way in which a bacteria strand becomes resistant is usually a mutation in a chromosomal gene of the pathogen. Whiles a organism is being treated by specific antibiotics, the antibiotics will have an effect on 99.99% bacteria but not the bacteria that have undergone mutation that prevents a certain antibiotic having an effect on these bacterial strands will reproduce and by the theory of natural selection predicts that under these circumstances, the fraction of the bacterial population carrying genes for antibiotic resistance will increase. For example, a mutation in one gene may stop or reduce the pathogen’s ability to transport a particular antibiotic into the cell. (Jane B. Reece).
There are quite a few practices effecting as to how bacteria strands become resistant to antibiotics. One would be the unnecessary prescribing of antibiotics from doctors to patients, but there is a lot to be said about this whether it be the patient feeling they are too sick to be told that they do not require the use of antibiotics even though they could just have a viral infection which antibiotics would be of no use to them unless it were to relief their pain or that they just want their moneys worth in antibiotics.
Another cause to do with the doctor patient relationship would be the fact that the doctor would be unsure of what to prescribe if need be or just how much to prescribe!
There is also this looming fear for doctors that the patient might wish to make a lawsuit against them for not taking action on their symptoms or not prescribing the right medication to them first time round and therefore doctors can be prescribing antibiotics out of fear of lawsuit.
Also, many practitioners who earn by means of selling medicines often prescribe more drugs than necessary for means of profit (Holloway 2000).
Another major factor that promotes bacteria to become antibiotic resistant is that when people do get prescribed the right medicine or antibiotics is that they don’t take the right amount each day. Some believe that it is better to take one antibiotic a day rather than two (Kardas P, March 2007) and others feel that it is ok to stop taking them when their symptoms have gone or that they will save them for the next time those symptoms occur. Its funny to actually hear that a third of people still believe that antibiotics are effective on the common cold (McNulty CA et al, August 2007).
In hospitals, poor hygiene can be associated with the contraction of noscomial infections and increase the risk of substaining a resistant microorganism, one of these well known noscomial infections is MRSA. Medical staff in hospitals world wide have been urged to wash their hands inbetween viewing patients and not to wear jewlery like wedding rings, bracelets or chains of the sort as these can transmit the infection from person to person (Girou E, Legrand P, Soing-Altrach S, et al October 2006). Much has been done in hospitals to stop the spread of noscomial infections but the treat still lingers with a massive one in seven chance of picking up a noscomial infection.
Another factor as to how we can contract resistant bacteria is by the food we eat.
Farmers feed their livestock antibiotics for numerous reasons but the fact is if and when their livestock build resistance to the antibiotics, they are then killed and processed into meats and other sources of food and they become our food. They may tell you your daily requirements for calories, vitamins, calcium, iron etc. but they do not tell you that your food could be the source of your illness or the reason why certain antibiotics will not have an effect on you!
 

Bacterial Magnetosomes Synthesized by Magnetotactic Bacteria Essay

ABSTRACT:
Bacterial magnetosomes synthesized by magnetotactic bacteria have recently drawn immense attention due to its unique features. Immobilized enzymes have a number of applications in today’s industries. Studies have shown that immobilized enzymes have a better shelf life and kinetics when compared to free enzymes. Magnetosomes have been used experimentally as carriers for antibodies, enzymes, ligands, nucleic acids, and chemotherapeutic drugs. This study reports the efficient immobilization of the enzyme alpha glucosidase on the magnetosomes extracted from the strain Magnetospirillum gryphiswaldense (MSR1). The alpha glucosidase enzyme was immobilized onto the extracted magnetosome surface using p-NPG (4-nitrophenyl-α-D-glucopyranoside) as the substrate. Theactivity of the magnetosomes was characterized by SEM analysis. There was successful immobilization of the enzyme on the magnetosome. It was observed that the properties of the immobilized enzyme had improved when compared with free enzyme.
Keywords: Magentotactic Bacteria, Magentosomes, Immobilization, Alpha glocosidase, Enzyme Activity.
INTRODUCTION:
Alpha-glucosidase (α-D-Glucosideglucohydrolase) is also known by another commonly used name, maltase. It plays a role as a catalyst in the hydrolysis of maltose to glucose units. Mammalian intestinal mucosa is known to secrete disaccharidases such as maltose, lactose and sucrose in abundance. Alpha-glucosidase (maltase) is used for inspecting the activity of alpha amylase and also for the determination of maltose in brewing. [6,7] Alpha-glucosidase catalyzes the following reaction;
α-GLUCOSIDASE
α-D-Glucoside + H2O → D-Glucose + Alcohol
Alpha-glucosidase hydrolyzes terminal non-reducing 1-4 linked alpha- glucose residues to release a single alpha- glucose molecule. Alpha-glucosidase is a carbohydrate-hydrolase that releases alpha-glucose as opposed to beta-glucose. Beta-glucose residues, on the other hand, can be released by glucoamylase, a functionally similar enzyme. The substrate selectivity of alpha-glucosidase is due to sub-site affinities of the enzyme’s active site. Two proposed mechanisms include a nucleophilic displacement and an oxo-carbenium ion intermediate.
Deficiency of α-glucosidase has ill effects on the insulin uptake system in the body causing a disease called Pompe’s Disease. Diagnosis of azoospermia has potential to be aided by measurement of alpha-glucosidase activity in seminal plasma. Inhibition of alpha-glucosidase can prevent fusion of HIV and secretion of HBV.
Clearly, this enzyme is very important both physiologically and medicinally or pharmaceutically. Immobilization of this enzyme is the main objective of this study. By immobilizing α-glucosidase onto the magnetic nanoparticles (magnetosomes in this case) the shelf life of the enzyme, its optimum temperature and optimum pH requirements and hence its storage and activity can be enhanced. This Alpha-glucosidase immobilization can be used as biopolymer and biomaterials in the field of pharmacy and medicine. [6,7]

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Magnetotactic bacteria (MTB) are phylogenetically diverse microorganisms that navigate along earth’s magnetic field. Most of these bacteria are microaerophiles that require a lower level of oxygen than what is provided at normal sea level on earth. High numbers of MTB (105-106/ml) are usually found at the oxic-anoxic transition zone, generally located at the sediment-water interface. Although MTB occur ubiquitously in diverse aquatic environments, different habitats have been found to contain various morphological types of MTB, including rod-shaped, vibrio-like, coccoid, and helical forms. A great diversity is also noted in the number, arrangement and shape of magnetosome particles in different MTB.
MTB occur in different phylogenetic lineages of bacteria. The majority of MTB affiliate with the α-Proteobacteria. A morphologically distinct, large magnetic rod was assigned to the Nitrospiraphylum, whereas a magnetotactic many-celled prokaryote and a magnetic sulphate-reducing bacterium belong to the δ-subclass of Proteobacteria. [8]
Magnetospirillum gryphiswaldense (MSR1) is a Gram-negative, spiral shaped, aquatic, mesophillic organism. Its optimum temperatures range is between 25-40°C. The main structures of interest inM.gryphiswaldenseare called the magnetosomes. Magnetosomes are the key component of these magnetotactic bacteria. They are intracellular, membrane-bound magnetic iron-bearing inorganic crystal like structures. It has been realized that these bacterial magnetosome portraya genuine prokaryotic organelle, showing a similar degree of complexity as observed in its eukaryotic counterpart. The morphology, composition, size and arrangement of these magnetic mineral crystals are subject to a species-specific chemical, biochemical and genetic control.Mature magnetite crystals usually fall within the size range of about 35–120nm. Exception being the very large magnetite crystals with lengths up to 250nm produced by an uncultured coccus.
The morphology of magnetosome produced usually varies between different bacterial species or strains. There are generally cuboidal, elongated or shaped like an arrow. Magnetosome are normally 35–120 nm long. Hence, single-domain. Magnetosomes are usually arranged in chains in the bacterium.[9]
Due to the magnetic interactions between the magnetosome which are arranged in chains, their magnetic dipole moments incline towards orientating themselves parallel to each other. The magnetic dipole moment of the bacteria overcomes the thermal forces while interacting with the earth’s magnetic field. These forces stray the position of the cell when in aqueous solutions.
The tiny magnetic dipoles in the magnetosome chain, when combinedfor working, make use of certain magnetic signals which help in the rotation of the bacterium towards the desired direction.Due to these forces, there is a twisting motion (torque) on the static magnetosome chain which will allow it to move the cell with the magnetic field lines. The magnetosome direct the bacteria towards a region where the environment for the growth of the cell is appropriate, but flagella are required for it to travel there.
MATERIALS AND METHODS:
Culturing and maintaining the organism:
All chemicals were purchased from Hi-media, India. TheHungateanaerobictechniquewas used with astandardprocedure for bacterial culturing and maintenance. Pure strain of Magnetospirillum gryphiswaldens (MSR 1) was obtained from DSMZ Germany. The culture was preserved in the standard condition as per the manual. The sub-culturing of the bacteria was carried out in the lab using MSGM medium. Modified MSGM medium was prepared by adding (v/v) of different percentages (5,10,25,50,75%)of nutrient broth (NB) (HIMEDIAM002) separately after adjusting MSGM media pH to 6.7 using NaOH.
70ml of the medium was measured and poured into serum bottles. Nitrogen was bubbled into the medium to create micro-aerobic condition. The bottles were closed with butyl rubber stoppers and sealed with aluminium caps. Media was sterilized by autoclaving and sterile solutions of wolfe’s vitamin elixir and ferric quinate was added. The media was then inoculated with Magnetospirillum gryphiswaldense culture and incubated in the shaker incubator for 3 days at room temperature.
Extraction of magnetosomes:
Three day old culture was transferred to 50ml centrifuge tubes and centrifuged at 8000rpm for 20 minutes. The pellet obtained was re-suspended in 10ml TRIS-HCl buffer (pH-7.4).(Tris HCl buffer was prepared by dissolving 0.5M Tris base or 61g of Trizma in 1000ml of distilled water).A 1:10 dilution of buffer and water was prepared before use. It was stored in a cool dry place. The suspended pellet was subjected to sonication for 120min (30W). After the sonication process, 1% SDS was added and incubated in water bath (90°C) for 5 hours. A black pellet was obtained.The tube containing the pellet and suspension had the magnetosomes. At every step the magnetosomes were washed using the buffer. The magnetosomes were separated by placing South Pole of a ceramic bar magnet near the magnetosome suspension. Magnetosomes were further purified by overnight incubation in Single tube magnet apparatus. The separated magnetosomes were lyophilized prior to immobilization of enzyme. [1]
Immobilization of α-glucosidase enzyme on magnetosomes:
Theα-glucosidase (167U/mg) used in this report was purchased from SLR, India. 4-nitrophenyl-α-D-glucopyranoside (p-NPG) was purchased from SLR, India. Glutaraldehyde (25% in v/v).
First, 100µL of 100 mmol phosphate-citric acid buffer (pH-5.0) containing 0.2% (v/v) gluraldehyde was added to 10µL magnetosomes (4mg/ml). It was subjected to ultrasonic dispersion for 20mins at room temperature. Then 1mL of α-glucosidase solution (0.05mg/mL) was added to the mixture and incubated at 25°C with shaking (150 rpm).Magentosomes with immobilized α-glucosidase were collected magnetically and washed thrice with 100mmol phosphate-citric acid buffer (pH 5.0). The activity recovery of the immobilized α-glucosidase was calculated as
Activity recovery (%) = [ Aimmob/ Ainit] X 100%
[Where Aimmob is the immobilized enzyme activity and Ainitis the initial (free enzyme) activity] [5]
The morphological and magnetic properties of the magnetic nanoparticles were characterized by SEM.
Determination of α-glucosidase activity:
α-glucosidase activity was determined by adding different concentrations of enzyme solution to 100mM phosphate buffer (pH-6.9) containing 10mM p-NPG. The reaction mixtures was incubated at 50°C in a water bath for 15mins and stopped by adding ml of 2M Na2CO3. Subsequently, the released p-nitrophenolwas measured at λ=405nm. [5] (One unit of α-glucosidase assay is defined as the amount of enzyme to release 1µmol of p-nitrophenol per minute.) Shelf life of the enzyme was determined by checking its kinetics one a week for 3 weeks.
RESULTS AND DISCUSSION:
After three months of culturing of Magnetospirillum gryphiswaldensein about 4 litres of MSGM, luxuriant growth was obtained containing about 10mg of magnetosomes. The magnetosomes were able to differentiate between the south and North Pole of a magnet and migrate. They responded only to the south pole of the ceramic magnet.This phenomenon is called magneto-taxis (fig. 1). If the bacteria obtained are a native of the northern hemisphere it is north-seeking. If the bacteria obtained are a native of the southern hemisphere then it is south-seeking. [11] The magnetosomes obtained in this case were south seeking.
Fig 1. Magnetosomes attracted to south pole of a magnet
SEM ANALYSIS: The morphology of the magnetosomes were analysed using scanning electron microscope and they were found to be of cubo-octahedral geometry. (Fig. 2a and Fig. 2b)

Fig 2a: Magnetosomes under scanning electron microscope

Fig 2b: Magnetosomes under scanning electron microscope
ENZYME ASSAY RESULTS:
To confirm immobilization, the activity of the enzyme-magnetosome complex was checked with that of the control (non-immobilized enzyme) at ÊŽmax=405 at pH 6.9 and temperature 29°C. A graph of % activity was plotted.

Table 1: % Activity of free enzyme and immobilized enzyme
The highest activity was observed to be 83.04%. This tells us that the activity is highest around pH=6.9 and temperature 29°C. (Table 1)
If pH and temperature was varied, the activity changed. [3]
The next assay performed had different concentrations of the enzyme combined with the same concentration of the substrate. The absorbance was checked at 405nm. % Activity recovery was calculated and a graph of % Activity versus concentration of the enzyme was plotted.

Table 2a: Acitivity with different enzyme concentration.
As the concentration of the immobilized enzyme increased, the % activity recovery also increased at pH 6.9 and temperature 29°C. (Table 2.)
The next assay performed had different concentration of substrates but same concentrations of enzyme. Again, the % activity was calculated and a graph of concentration versus % activity was plotted.

Table 2b:Activity with different substrate concentration
The % Activity increased with the increase in concentration of the substrate. The pH was 6.9 and temperature was 29°C.( Table 2b)
STORAGE STABILITY:
It was observed that the activity of the enzyme had not been altered even after storing it at 4°C for three weeks. The % activity were found to be 81%, 72% and 72% after 1,2, and 3 weeks respectively.
CONCLUSION
It has been observed on the whole that the activity of α- glucosidase was enhanced when immobilized onto the magnetosomes.This activity however has not been reported before. Immobilization onto magnetic nanoparticles has been reported but using magnetosomes this is one of the only papers that has successfully immobilized α- glucosidase and demonstrated enhanced activity in terms of %activity.