Antibacterial resistance in bacteria has become an apparent problem in the world today. This is due to the fact that antibiotics have been prescribed and used too often in our daily lives. This is unfortunate for medicine because all antibiotics, including the most effective, vancomycin, are now at risk of failing to protect us from harmful bacterial diseases. Antibacterial resistance is believed to take place more often in areas where antibiotics are frequently used. This study was conducted to test for more antibiotic resistant bacteria found in a shower frequently cleaned with 'Spic-N-Span' versus a shower not often cleaned with antibiotic products. This study provides some evidence for antibacterial resistance in bacteria present in the 'Spic-N-Span' shower environment. A few key methods used to test this experiment included the following: the Gram stain test to determine whether or not the bacteria was Gram-negative or Gram-positive, agarose gel electrophoresis to visualize the DNA to compare it to a known base pair sample, restriction enzyme digest test to determine where our DNA plasmid cuts, transformation to transform competent E. coli cells, and a PCR test to replicate large quantities of a targeted region of DNA. While all of the results from our experiments initially supported our hypothesis that the 'Spic-N-Span' shower produces more antibiotic resistant bacteria than the dirty shower, statistical tests showed that the environments were not significantly different. The "Red" control our team used was resistant to ampicillin. We concluded that overusing antibiotics in the same environment could eventually lead to antibiotic resistant strains of bacteria. On a larger scale, this is unfortunate news for the world we live in. Antibiotic resistant bacterial strains can develop quickly and spread even quicker. The results obtained from researching different types of bacteria in varying shower environments can be put to practical in scientific studies trying to determine how to stop or limit antibiotic resistant bacteria. Further experimental studies that could be conducted might include trying to determine another sanitizing product that does not involve antibiotics, but still gets the job done and the desired environment clean.
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Antibiotic Resistance is becoming an increasing problem in today's society. There has currently been a decrease of useful antibiotics on the market due to the bacteria's increasing resistance to antibiotics (Rambhia and Gronvall, 2009). Antibiotics are one of the leading ways to treat potentially life threatening bacterial infections. Due to their overuse, many bacteria have become resistant to antibiotics.
Most of the research conducted in the field has been done on showerheads and hospitals. Hospitals are particularly prone to resistance because of the vast amounts of bacteria being treated with antibiotics on a daily basis. In addition, there are places that these potentially more resistant bacteria harbor and can then multiply and infect more patients. One such study was about Pseudomonas aeruginosa, a strain of bacteria that can lead to severe infections. P. aeruginosa is a strain that is generally only found in immunocompromised patients. The resulting infection was increasing in hospitals and eventually traced back to the type of bathtub used. The bathtubs had a certain type of strainer on the drains that could harbor bacteria. When the bathtub was filled, the P. aeruginosa would spread throughout the entire bathtub and greatly increase infection risk of the person in the bath. This was especially true if that person had open cuts and/or a compromised immune system, both of which are common cases in a hospital setting (Berrouane et al., 2000).
Showerheads can also be linked to the spread pathogenic bacteria. The insides of showerheads are an ideal ecological niche for colonizing bacteria. The showerhead provides a warm, moist, dark place for potentially harmful bacteria to grow. In addition to potentially infecting a person who is directly under the initial water blast, the shower can aerosolize microorganisms, which can increase pulmonary risk (Feazel et al., 2009).
In an experiment conducted by Kohanski et al. (2010), antibiotics, such as ampicillin and kanamycin, were placed on culture plates to test the mutation rates of a specific E. coli strain. The author's overall objective was to determine how much, if any, mutation would occur in the specific E. coli strain and how much antibiotic resistance might result. Samples of five different antibiotics: norfloxacin, ampicillin, kanamycin, tetracycline, and chloramphenicol, were taken and treated for five days in two separate environments. In one environment, the ampicillin treatment was genetically crossed with the wild-type E. Coli to see if any antibiotic resistance would occur. After comparing the results, the bacterial samples that were treated with ampicillin had a greater percentage increases in the amount of antibiotic resistant bacteria they contained. This is in contrast with the low percentage increases produced by the no drug treatment (Kohanski et al., 2010). Although this experiment did not involve bacteria from a shower-like environment, the same antibiotics were used to test a certain strain of bacteria and how it would mutate if it were genetically crossed with other antibiotics.
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Antibiotics can work in a variety of ways. One way is by binding to ribosomes and inhibiting protein synthesis. Kanamycin and tetracycline are two common antibiotics that use this method. Antibiotics such as ampicillin and doripenem work by inhibiting the enzyme involved in creating the bacterial cell wall. Different antibiotics can also inhibit synthesis of DNA, tetrahydrofolic acid, or block active sites in the protein. All of those will eventually destroy a population of non-resistant bacteria. (Alkeshun et al. 2007)
Despite the effectiveness of antibiotics, bacteria are not helpless. Due to rapid reproduction, mutations, and conjugation, bacteria can become resistant to antibiotics. On a molecular level, bacteria can become resistant by creating enzymes that degrade or inactivate antibiotics. Some bacteria replace the proteins or enzymes affected by the antibiotics. Bacteria can also modify themselves to prevent entry by the antibiotic, or create a kind of pump that will just pump the antibiotic right back out of the cell (Levy, 1998). Bacteria can share any mutations with other bacteria by a process called horizontal conjugation. Horizontal conjugation occurs when a plasmid within a host organism pulls up along- side another plasmid and cuts into the host plasmid's DNA sequence and inserts its own genetic material (Alekshun et al., 2007). These tactics all have biological cost and reduce the fitness of the bacterial cell. If an antibiotic is not prevalent, then the non-resistant bacteria will eventually take over due to the resistant cells loss in fitness. This is one way that scientists hope to reduce antibiotic resistance (Andersson, 2003).
On a large scale, populations of resistant bacteria are usually formed by overuse or misuse of antibiotics. When non-resistant bacteria are killed by an antibiotic, any resistant bacteria will remain and multiply. Over time, the only populations of the bacteria remaining will be resistant. Sometimes bacteria are only insensitive to an antibiotic, meaning that it will take more exposure, but the antibiotic will still eventually kill the insensitive bacteria. If an antibiotic is not used for long enough, the insensitive bacteria will still be active and multiply, making all of the bacteria insensitive.
Our experiment analyzes the bacterial growth on the drains of two Holmes Residence Hall showers; one often cleaned with 'Spic-N-Span,' and the other considered dirty, due to its lack of prior cleaning. Both showers were swabbed and the bacteria was isolated onto master plates and then swabbed onto three antibiotic and control patch plates. These plates were analyzed for antibiotic resistance. This was determined by testing and observing the bacteria as they grew on three different antibiotic plates containing ampicillin, tetracycline, or kanamycin. If the bacteria still grew with the presence of the antibiotics, it showed that the antibiotic didn't work and the bacterium was resistant to the antibiotic. Once resistance was found, a colony of the patch plate was streaked onto an antibiotic streak plate. A single colony grown on these plates was then extracted into a liquid medium to be further analyzed.
We hypothesize that the drain that has been previously exposed to more antibacterial cleaner, the 'Spic -N-Span' shower, will have more bacteria resistant to the antibiotics. We predict that because the 'Spic-N-Span' shower has been introduced to antibacterial products, the bacteria within the environment will be more resistant to antibiotics such as kanamycin, tetracycline, and ampicillin. The independent variable being tested is the changing environments, both the dirty and 'Spic-N-Span' shower surfaces. The dependent variable is the number of bacterial colonies being grown on each of the antibacterial plates. The control is the Lysogeny Broth (LB) plates that were streaked last to ensure bacteria made it on to the antibiotic plates and had potential to grow. We found that the bacteria in the 'Spic-N-Span' grew more colonies on antibiotic plates than the Dirty bacterium did.
Acquiring Bacteria and Setting up LB Plates
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. In order to obtain the bacteria from the shower drain, a sterile swab dipped in phosphate-buffered saline (PBS) was used to scrape the inside of the drain for bacteria. The swab tips were then streaked onto Lysogeny Broth (LB) agar plates. One must be careful to not pass the swab over the same area more than once; this ensures the amount of bacterial growth is not too excessive to be able to distinguish between individual colonies of bacteria. The plates were then incubated for 24 hours at 37Â°C. After the bacteria on the LB swab plates have grown for 24 hours, they were sealed with parafilm and refrigerated.
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Master LB Plates
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. A master LB plate was created by dividing a standard LB agar plate into 16 squares and using a sterile loop to streak a separate singular bacterial colony into each of the 16 squares. A unique colony was chosen from the LB streak plate and a one centimeter streak was made inside a single square of the master LB agar plate. This process was repeated for all 16 squares of the master patch plates. The master plates were then incubated for 24 hours at 37ËšC, sealed with parafilm, and refrigerated.
Master Patch Plates
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. These plates differ from the master LB agar patch plates because the antibiotics ampicillin, tetracycline, and kanamycin, were added to the agar in these plates. These antibiotic plates were also divided into 16 squares in which a streak of square number one of the master streak plate was transferred to square number one of the three different antibiotic plates and an LB agar plate was used as a control. It was important that the antibiotic plates were streaked in the same order, making the LB agar plate last as a control. The order of the plates was: ampicillin, kanamycin, tetracycline, and LB agar.
Streaking of Individual Colonies
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of streaking the individual colonies on antibiotic plates was to isolate well-defined, individual colonies. A specific, well-defined colony was chosen from the master patch plate and isolated onto an LB only plate by the use of a cool, sterile loop. The bacterium was spread back and forth several times using the loop, covering about one-fourth of the plate, known as the first quadrant. The loop was re-sterilized and cooled, and then placed in the midst of the last few streaks of the first quadrant and was then used to spread the bacteria around the plate in the second quadrant. The same procedure was repeated for quadrants three and four. The streak plates were then placed in the incubator for twenty-four hours at 37ËšC.
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of a KOH test was used to determine the type of cell wall, either Gram-positive or Gram-negative, of the bacterium and where it is located. To begin, 10-20 Î¼L of 3% KOH was placed onto a microscope slide. Bacterial cells were then added to the slide with a cooled, sterile loop and swirled for approximately one to two minutes. Thickening signifies that the slide contains Gram-negative bacteria. If the liquid did not thicken, it signified that it contains Gram-positive bacteria. If this is the case, add 250 Î¼L of cell suspension solution and 5 Î¼L of 50 mg/mL of stock solution. It was incubated for thirty minutes at 37ËšC, and the cell lysis step was then repeated.
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of the Gram stain test was to identify if the bacterium is Gram-positive or Gram-negative based on the chemical and physical properties of their cell walls. To start, a colony of bacteria was smeared onto a microscope slide and was allowed to dry. The slide was then moved over the flame of a Bunsen burner to affix the bacteria to the glass. The entire slide was then flooded with crystal violet and was then allowed to stand for about sixty seconds, and then rinsed for five seconds with water. The specimen should have appeared blue-violet when observed with the naked eye. The slide was then flooded with iodine solution and was allowed to stand for about a minute. The slide was then rinsed with water for about five seconds and was immediately passed on to the next step. The slide should also still have appeared blue-violet at this point. Ethanol, the decolorizer, was then added drop-wise until the blue-violet was no longer being emitted from the specimen. The ethanol was added drop-wise to be safe because adding too much decolorizer could result in a false Gram-negative result, whereas not using enough decolorizer may result in a false Gram-positive result. After the addition of ethanol, the slide was rinsed with water for five seconds. The slide was then flooded with safranin and allowed to stand for about one minute to allow the bacteria to incorporate the safranin. Gram-positive cells were determined with the blue-violet appearance and Gram-negative cells were determined with a pink color appearance. The slide was then rinsed with water for five seconds to remove any excess dye. After this was done, the slide was wiped gently with a Kimwipe and allowed to air dry before viewing it under the microscope. A small amount of immersion oil was placed between the immersion lens and the slide and then the lens was moved down to meet the oil. The team made sure that the 100X lens was being used during this process.
Liquid Media Culture and Bacterium
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The liquid media bacterial culture and the bacterium were used for plasmid isolation by the Promega miniprep protocol. It is important to note that if your bacterium is Gram-positive, you may have trouble lysing cells. If this occurs, take 5 Î¼L of the 50 mg/ml stock lysozyme and mix with 250 Î¼L of cell suspension and incubate at 37ËšC for thirty minutes. Then you can proceed to the starting step. To start, 5 mL tubes with liquid medium were obtained. 5 Î¼L of appropriate antibiotic ampicillin, kanamycin, or tetracycline, was added to the liquid medium. A single colony from an antibiotic streak plate was sampled with a sterilized loop and dipped into the medium and mixed together. The tube was loosely capped and placed in the shaker for twenty-four hours.
Plasmid Miniprep Protocol
The purpose of a Wizard Plus SV Minipreps DNA Purification System plasmid miniprep was for the isolation of a plasmid for determination of plasmid-mediated antibiotic resistance (Promega, Madison, WI). To start, 1-10 mL of overnight liquid medium cultures were pelleted for five minutes. The pellets were then thoroughly resuspended with 250 Î¼L of cell resuspension solution. 250 Î¼L of cell lysis solution was added to each sample and then inverted four times to mix. 10 Î¼L of alkaline protease solution were added and then inverted four times to mix. The pellets were then incubated for five minutes at room temperature. 350 Î¼L of neutralization solution was then added and inverted four times to mix. The pellets were then centrifuged at top speed for ten minutes at room temperature. A spin column was then inserted into each collection tube. Cleared lysate was then decanted into the spin column. The collection tubes were centrifuged at top speed for one minute at room temperature. Flowthrough was discarded and the spin column was reinserted into the collection tube. 750 Î¼L of wash solution was added to the collection tubes and centrifuged at top speed for one minute. Flowthrough was discarded and the spin column was reinserted into the collection tube. This step was repeated with 250 Î¼L of wash solution and then centrifuged at top speed for two minutes at room temperature. The spin column was transferred to a sterile 1.5 mL microcentrifuge tube to ensure that none of the column wash solution was transferred with the spin column. If the spin column still had column wash solution associated with it, it was centrifuged again for one minute at top speed, and then transferred to a new sterile 1.5 mL microcentrifuge tube. 50 Î¼L of nuclease-free water was added to the spin column and centrifuged at top speed for one minute at room temperature. The spin column was discarded and the DNA was stored in a collection tube at -20ËšC.
Agarose Gel Electrophoresis
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. To begin, a 1% agarose gel was made by melting 0.4 grams of agarose in 40 mL of 1X concentration of TBE buffer. The melting was done by use of a microwave and the team made sure that the solution did not boil over. After the solution was cooled to about 50-60ËšC, 2 Î¼L of 1 mg/mL of Ethydium Bromide (EtBr) was added to the solution. The entire was solution was then poured into a gel mold. Gloves were worn the entire time during the process of handling EtBr. The DNA sample was prepared next. To do this, 10 Î¼L of plasmid DNA from the miniprep was placed in a tube with 2 Î¼L of 10X concentration gel loading buffer. After the agarose gel cooled and solidified, it was loaded with DNA ladder and environmental and control plasmid samples. The gel was then exposed to an electric field and was run for approximately thirty to sixty minutes at 100 volts. After that time had elapsed, the gel was visualized by ultraviolet illumination using a Kodak imaging system.
MacConkey Agar Test
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The MacConkey agar test was utilized to grow Gram-negative bacteria and stain them for lactose fermentation. To start, a streak plate was made on MacConkey agar and then incubated for two to twenty-four hours at 37ËšC. After the incubation, the plates were analyzed to determine which plates contained bacterial growth, if any. If the growth was yellow, that signified that the bacteria could not ferment lactose. If the growth was pink, that signified that the bacteria could ferment lactose.
Eosin Methylene Blue Plates Test
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of the EMB plates was to inhibit the growth of the Gram-positive bacteria to be used as another test for Gram identity. The procedure for this experiment is the same as the MacConkey Agar Test (see above). If the plates contained bacterial growth, this implied that the bacteria was Gram-negative.
Restriction Endonuclease Digest Test
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of the restriction endonuclease, or enzyme, digest test was to cut double or single stranded DNA at specific recognition nucleotide sites called restriction sites. The resulting fragments can be measured and compared to a known plasmid to confirm identity of the experimental plasmid. In preparation for a restriction enzyme digest, a 20 Î¼L reaction was required, including 10 Î¼L of DNA, 2 Î¼L of enzyme (one for each of the two enzymes, EcoRI and BamHI), 2 Î¼L correct buffer, 1 Î¼L BSA, and 5 Î¼L of water. This mixture was then incubated for approximately twenty hours at 37ËšC. 10Î¼L of this solution was taken and placed into a gel along with 10 Î¼L of "Red" control DNA and 2 Î¼L of 1 kb Invitrogen ladder to test where the enzymes will cut.
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of this procedure was to transform competent E. coli cells. To start, one 50 Î¼L aliquot of competent cells was obtained and thawed on ice. 22 Î¼L of the cells were pipetted from one tube into a new 1.5 mL microcentrifuge tube. 1 to 5 Î¼L of "Red" plasmid DNA was added into a vial of competent cells and stirred gently with the pipette tip to mix. A control containing no plasmid was made by adding 1 to 5 Î¼L of H2O into a separate tube of competent cells and was mixed gently. The tubes of cells were then incubated on ice for thirty minutes. The cells were then heat-shocked for 30-45 seconds at 42ËšC without shaking. The vials were removed from the 42ËšC bath and placed on ice for two minutes. 250 Î¼L of pre-warmed (37ËšC) SOC medium was added to each tube of cells. The tubes were tightly capped and shaken horizontally at 37ËšC for one hour at 225 rpm in a shaking incubator. 75 Î¼L of each transformation were spread, using a "hockey stick," onto a pre-warmed "LB only" plate and 75 Î¼L were spread onto both "selective" LB/Amp and LB/Kan plates. The remaining transformation mix was stored at 4ËšC in a refrigerator. The plates were inverted and then incubated overnight at 37ËšC. Each of the transformation plates was photographed and the number of colonies growing on each one was recorded. If the plates had too many colonies to count, it was indicated as a "lawn."
Polymerase Chain Reaction Test
The following protocol was obtained from LB145 Spring 2010 Lab manual constructed by the LBC Biology Staff. The purpose of the PCR test was used to replicate, or "amplify", large quantities of a targeted region of DNA in vitro. To start, all of the following reagents were added into one microfuge tube: 80 Î¼L Nuclease-free water, 10 Î¼L Thermopol buffer, 3 Î¼L dNTPs, 2 Î¼L primer 11F, and 2 Î¼L primer 1492R. 1 Î¼L of Taq polymerase was added to the master cocktail, making the concoction 98 Î¼L total. The microfuge tube was vortexed for 1-2 seconds two or three times. 30 Î¼L of the master cocktail was pipetted into each of the three microtiter reaction tubes (PCR tubes): one environmental bacteria, one E. coli, and one negative control (water). 10 Î¼L of SOC medium was taken and had one bacterial colony added to it, vortexed for 1-2 seconds, and then 1 Î¼L of the resulting solution was added to the PCR tubes. All of the PCR tubes were then labeled. A micropipette with a tip was then used to transfer one colony of E. coli from the plate into an appropriately labeled PCR tube. This step was repeated using the environmental bacterium instead of E. coli. 1 Î¼L of water was then added to the remaining PCR tube. The PCR tubes were then placed into the thermal cycler. The tubes initially underwent initial denaturation for five minutes at 94ËšC. The tubes then endured denaturation for 30 seconds at 94ËšC, annealing for 30 seconds at 50ËšC, and extension for 45 seconds at 72ËšC. This step was repeated for 35 cycles. The PCR tubes underwent a final extension for seven minutes at 72ËšC. The LAs then removed the PCR tubes from the thermal cycler and placed them in the freezer until our team was able to come to lab and run a gel on them. To run the gel, 10 Î¼L of each reaction mix, plus 2 Î¼L of loading dye, were placed into an agarose gel, just like for the plasmid miniprep and restriction enzyme digests. 2 Î¼L of 1 kb ladder from Invitrogen was also added into the gel. The gel was run for approximately 30 minutes and was then photographed. Further steps included the clean-up of PCR products in preparation for 16S sequencing and analysis of the purified PCR product and sample preparation for sequencing.
Master Patch Plates
The number of colonies on each antibiotic plate varied based on the antibiotics used. After the previous shower samples (Figure 1) were streaked onto the antibiotic plates, tetracycline was shown to contain five total colonies for the three dirty shower trials and a total of eight colonies on the 'Spic-N-Span' shower plates. Ampicillin had 47 (out of a maximum of 48) colonies on the 'Spic-N-Span' plates, and only 31 for the dirty shower samples. The kanamycin plates had 42 colonies for the dirty shower and 45 for the 'Spic-N-Span' environment. Bacteria grew on all sixteen colonies on every LB plate, regardless of the environment or trial number (Figure 2). In all cases, the 'Spic-N-Span' shower had more colonies grown than the dirty shower (Table 1). The patch plates chosen for re-streaking onto another antibiotic plate to be further analyzed were: The first and second trials of the 'Spic-N-Span' kanamycin plates, both squares 7 ('Spic-N-Span' #1 kanamycin plate); the second trial of the dirty kanamycin plates, square 5 (dirty #2 kanamycin plate); and the third trial of the dirty ampicillin plates, square 2 (dirty #3 ampicillin plate) (Figure 3).
We were unsuccessful in isolating a plasmid from our shower samples, however, we did isolate the "Red" control sample. This DNA was used for most of the future experiments for this stream (Figure 4).
When the KOH test was applied separately to the bacteria from each of the four antibiotic sample streak plates, the solutions thickened, showing that the bacteria from all of the antibiotic plates were Gram-negative (Figure 5). Gram stain tests were performed on the bacteria from the four sample streak plates, all appeared pink, thus holding the saffron dye, indicating that the bacteria sampled were Gram negative (Figure 5). In the MacConkey Agar tests, two of the plates did not have any growth, indicating that the bacteria were Gram positive. 'Spic-N-Span' #1 kanamycin plate and dirty #3 ampicillin plate both came up Gram negative while the other two appeared Gram positive (Figure 6). Last, an EMB test was performed. It was similar to the MacConkey agar tests. Bacteria grew on all of the EMB plates, indicating that they are all gram negative (Figure 7) (Table 2).
Restriction Endonuclease Digest Test
We were unable to successfully complete the restriction enzyme digest. After numerous trials using different enzymes each time, the enzymes never successfully cut the plasmid (Figure 8).
Both LB plates for the "Red" control bacteria and the water had lawns of bacteria on them. Neither antibiotic had anything grow on them when water was used. This was expected because the water has no resistance to ampicillin or kanamycin. If it did have resistant DNA, the E.coli would be able to transform resistance against that antibiotic. For the "Red" control, the ampicillin antibiotic plate had eleven colonies, while the kanamycin had none. It was expected that colonies would grow on one of these plates but not the other because it was known that the bacteria used was resistant to either ampicillin or kanamycin, but to which one was unknown (Figure 9).
Polymerase Chain Reaction
The PCR test was also unsuccessful for the environmental tests. However, the E.coli was successfully replicated with the PCR. The gel electrophoresis showed a band at1636 base pairs, signifying that the plasmid was cut at 1636 base pairs and created one long fragment. (Figure 10). Because we already know what kind of bacteria the control E.coli is, there was little point in going through the clean up and sequencing it.
When calculating statistics using Vassarstats, the P-value was calculated to be 0.176, 0.823, and 0.434 for the ampicillin, kanamycin, and tetracycline, respectively. The Chi-square tests resulted in observed values of 1.83, 0.05, and 0.61 for ampicillin, kanamycin, and tetracycline, respectively. If the p-value is less than 0.05, then the null hypothesis is rejected. If the observed value of Chi-square test is greater than the actual Chi-square value from the Chi-square table (in this case 6.63), then the null hypothesis is rejected. In both cases, the statistics show that the null is not able to be rejected, so our hypothesis is unsupported.
The purpose of this experiment was to investigate a potential cause of antibiotic resistance in bacteria. The original theory of the experiment was to find evidence to support our idea that antibiotic resistant bacteria are more commonly found in environments where antibiotics are used more frequently. In our experiment we found that the shower more often cleaned with Spic-n-Span had more growth of antibiotic resistant bacteria. This supports our hypothesis and prediction, which stated that the 'Spic-N-Span' shower would contain more antibiotic resistant bacteria because of its prior exposure antibiotic products. However, the statistics did not show support that the environments were significantly different. The p-value was greater than .05 and the actual Chi-square value was greater than the observed value. This shows that we cannot support our hypothesis.
Methods such as the KOH test, Gram stain, EMB test, and two of the four of the MacConkey agar test all gave results that the bacteria were Gram negative. The other two MacConkey tests gave results of Gram positive bacteria (Figure 6). Gram identity tests are performed to discover which type of cell wall each bacterial cell contains. This is significant because this will determine whether the bacterial cell is Gram positive or Gram negative. This is important because a thicker cell wall, implying that it is Gram positive, means that antibiotics will have a harder time fighting off bacteria because it is harder to penetrate the cell. The inconsistencies in our Gram identity experiments could be due to human error in streaking or preparing streak plates or contamination of Gram positive bacteria, giving false results.
Our transformation results provided evidence that the control bacteria was resistant to ampicillin because there was bacterial growth on the ampicillin plate. The ampicillin plate contained eleven distinct bacterial growth colonies which serve as the basis of our support for our conclusion that the "Red" control was pAMP and resistant to ampicillin. The restriction enzyme digest also could have helped show that the "Red" control is pAMP if it had been successful. The enzymes should have cut the plasmid at specific base pairs. When the plasmid was compared to the 1 kb invitrogen ladder, it should have shown what the base pair lengths were, which we could have compared to the known plasmids of pAMP and pKAN to determine which antibiotic the bacteria was resistant to. Five unsuccessful restriction enzyme tests were performed in total; however, we do know that the "Red" control is pAMP due to the successful transformation. The restriction enzyme digest may have failed to work correctly due to human error in the restriction enzyme digest procedure, such as not having enough enzyme or using incorrect proportions of materials. Another way error could have occurred is if DNA was not isolated properly during the miniprep stage or the enzymes could have been faulty due to improper isolation or storage.
The PCR test was also unsuccessful for our environmental plasmids. This could have been due to poor technique in preparation of the reaction cocktails or individual materials for the cocktails, or an error occurring in the thermal cycler. The E. coli control was successfully replicated in the PCR with the plasmids being cut at 1636 base pairs. However, because E. coli is already a known strain, the clean up and sequencing of the bacteria was not necessary.
Despite the fact that the statistical data did not support our hypothesis, we can conclude that antibiotic resistant bacteria do exist more commonly in frequently used antibacterial environments due to previous findings such as Buckwold and Ronald (1979), Kohanski et al. (2010), Berrouane et al. (2000), Levy (1998) and Rambhia and Gronvall (2009) and the fact that all of our antibiotic exposed environments did contain more antibiotic resistant bacteria (Table 1). Human error could have occurred in obtaining the samples due to poor swabbing technique of the two shower environments or in our statistical calculation of the colonies. Statistical anomalies should not be disregarded; however, we feel that the previous research in the matter supports our original hypothesis more than our statistical results. What does this mean for medicine? These results can be used to help promote responsible antibacterial use to help maintain and control bacterial strains in the hope that they will not become any more resistant to antibiotics than they already have. Further experiments can be conducted in this area with the use of different antibiotics in similar types of environments as well as different types of environments for further comparison. Also, the same types of antibiotics can be used in different types of environments for further comparison as well. All experiments should be completed with the intent to provide results that may help medicine improve fighting off harmful bacterial diseases.