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Video Abstract:
Video Abstract:
AAQ Final Video Abstract .mp4Background: Biofilms / Bacteria vs Antibiotics
Background: Biofilms / Bacteria vs Antibiotics
Biofilms are dense, organized structures of microbes that grow on almost all surfaces: rocks, pond surfaces, banana skin, and even teeth. Consisting of four stages—attachment, biofilm formation, dispersal, and recolonization—biofilms’ life cycle begins when a specific environmental signal induces free-living planktonic cells to attach to nearby surfaces by means of flagella and other cell-surface appendages. Those attached cells coat the surface with an organic monolayer of polysaccharides or glycoproteins, to which other cells attach. During the biofilm formation stage, bacteria lose flagella and eventually stop to bind to the surface. As more cells bind and divide, they form microcolonies and communicate by sending and receiving chemical signaling molecules through quorum sensing (the regulation of gene expression according to population density). During the final life cycle stages of dispersal and recolonization, nutrients become limiting and cells leaving the biofilm revert to their flagellated, planktonic forms, moving to new locations with greater nutrient access. Finally, recolonization restarts attachment, and the process restarts.
Biofilms benefit bacteria through a few mechanisms. First, biofilms’ circulatory channels provide microbes with a stable microenvironment that gives them access to a nutrient source. Second, because microbes within a biofilm are packed in such close proximity to one another, they can communicate, behave cooperatively or competitively, and exchange genetic information. Finally, the extracellular polymeric substance (EPS) matrix that forms and surrounds a biofilm protects microbes from external threats and environmental stressors such as desiccation, predators, and host immune cells.
Biofilms can also tolerate antibiotics. Biofilms frequently grow on the surfaces of medical devices (MRIs and x-ray machines), implants (catheters and pacemakers), and dead tissues and contribute to the recurrence of chronic infections. This poses a serious threat, as roughly 80% of microbial infections are caused by biofilms. This resistance is in large part due to biofilms’ multicellular nature, as it allows for the delay and/or prevention of antibiotic penetration and diffusion into the extracellular matrix and altered chemical environments within the biofilm. Biofilms can also lead to the degradation of antibiotics through enzymatic reactions, horizontal gene transfer, target site mutations, efflux pumps, and drug-neutralizing proteins. Also, cells that have entered a stationary phase within the biofilm can become tolerant to antibiotics, leading to higher resistance among older biofilms.
Bacteria divide every 4 to 20 minutes, reproducing rapidly and most often through binary fission. In binary fission, a bacterium provides each offspring with a complete copy of its essential genetic material, so the genes and mutations present in the parent cell’s DNA will, barring a rare mutation, be present in the offsprings’ as well. In terms of the genes responsible for bacterial biofilm formation, then, the offspring of biofilm-forming parent bacteria will most always display the capacity to form biofilms as well. Among bacteria with different parentages, though, genetic disposition for biofilm formation varies. This variation makes natural selection, the process by which organisms (like bacteria) better genetically suited to their environment survive and reproduce at a higher rate than organisms that are not as genetically suited.
Why Bacteria, Antibiotics, and Biofilms?
Why Bacteria, Antibiotics, and Biofilms?
(Full written description linked here!!)
P. fluorescens and Amikacin
P. fluorescens and Amikacin
We selected P. fluorescens because it was the bacteria tested in the first biofilm lab/preliminary testing. Amikacin was chosen because the literature we consulted named amikacin as a potent antibacterial agent against a close relative of P. fluorescens, P. aeruginosa. These articles also cited a concentration threshold that determined amikacin’s potency, establishing both experimental signal and our antibiotic concentration: 1.06 μg amikacin per liter of water.
Biofilms' Antibiotic Resistance
Biofilms' Antibiotic Resistance
After finishing our first lab working with bacteria and biofilms, we decided to further investigate the dynamics between bacteria, selection for biofilm formation, and biofilms in general -- more specifically, their ability to protect bacteria from antibiotics. Antibiotic resistance poses a huge threat within the medical world, as bacteria can colonize and create biofilms on a huge range of surfaces like medical devices, implants, and living tissues.
Our results and the mechanisms behind them are key because they can inform research about the roles of biofilms in improving bacterial survival. If scientists and doctors seek to design antibiotics that infiltrate and destroy biofilms, they must understand the mechanisms those antibiotics have to overcome.
Articles:
Goal:
Goal:
To determine how the presence of biofilm formation selection among P. fluorescens affected the bacteria’s survival rate and resistance to an antibiotic, amikacin.
Hypothesis:
Hypothesis:
P. fluorescens receiving an amikacin injection on day 4 will survive at a higher rate than those receiving an injection on day 2 because the later-injected bacteria will have undergone the selection process and consist only of bacteria—and their offspring—that successfully form biofilms, and biofilms’ multicellular structure and extracellular matrix protect bacteria from antibiotic permeation.
Methods and Lab Setup:
Methods and Lab Setup:
Day 1: The antibiotic was made through the mixing of a concentration of 1.06 μg amikacin per liter of water. Four culture tubes were labeled: E1, C, E2, and C2 (C indicating the control group for the unselected bacteria and C2 the control group for the selected bacteria) and with our group name and the date. 4.5 mL of Queen's B Media (QB) solution was added to each of the four tubes with a serological pipette. A white polystyrene bead was inserted into culture tubes E2 and C2 with sterile forceps, and a single, isolated P. fluorescens colony was added to all tubes using a sterile inoculation tube. The tubes were then incubated on an orbital shaker until the next day.
Day 2: 45μl of the amikacin solution was injected into the Day 1 E1 culture tube. Two new tubes were labeled “E2” and “C2” as before, and 4.5 mL QB media was pipetted into each new culture tube before the tubes were set aside for later use. Separately, we added 950μL QB to four microcentrifuge tubes labeled “E1,” “C,” “E2,” and “C2,” all with the concentration marker 100. Three more groups of “E1,” “C,” “E2,” and “C2” tubes were labeled, except with concentration markers 10-1, 10-2, and 10-3, and all tubes were filled with 900μL of Phosphate Buffered Saline (PBS).
The group two, biofilm-forming bacteria was transferred: Day 1 beads from both the experimental and control cultures were moved with sterile forceps to the corresponding Day 2 culture tubes and vortexed for a minute to ensure that all bacteria detached from the beads. The Day 2 group 2 tubes were again incubated on an orbital shaker until the next day, and the unused E2 and C2 microcentrifuge were set aside.
Without transfers, Group 1’s bacteria culture samples were plated after a serial dilution process. Two Tsoy-Agar plates were labeled as "E1 10-2" and “C 10-2”, and another two were also labeled “E1 10-3” and “C 10-3.” For both control and experimental cultures, we transferred 50μL of the Day 1 cultures to the corresponding 100 microcentrifuge tube filled with 950 QB, and the tube was briefly vortexed to mix. 100μL was transferred from both 100 tubes to the corresponding 10-1 tubes, the tube was vortexed, and this step was repeated until both the 10-2 and 10-3 microcentrifuge tubes had been filled and vortexed. 100μL of the 10-2 and 10-3 dilutions were transferred to agar plates, spread evenly with a plate spreader (sterilized over a flame for each plate), and the plates were incubated upside down until the next day.
Day 3: group 2’s Day 2 bead was transferred in the same manner into the Day 3 tube and then vortexed. The plated group 1 samples were examined and photographed so that the number of surviving colonies could be counted manually or through ImageJ analysis.
Day 4: group 2’s beads were transferred for the final time into the Day 4 tube. 45μL amikacin was injected into the experimental culture before both the control and experimental cultures were diluted and plated in the same sequence as group 1, then incubated overnight.
Day 5: the group 2 plated colonies were photographed and counted, again manually or through ImageJ analysis.
Results
Results
Key Findings:
(Figure 4) P. fluorescens’ mean surviving colonies in both the control and experimental groups increased significantly (p<0.05) between the groups unselected and selected for biofilm formation
The unselected experimental group’s colony mean was 328±79
The selected experimental group’s mean was 6144±1248
Differences between the control and experimental group colony counts of both the unselected and selected bacteria were statistically insignificant (p=~0.463 and p=~0.648, respectively)
Key Findings:
(Figure 5) Mean survival rate was significantly greater (p=~3.239 x 10^-5) for selected P. fluorescens than for unselected P. fluorescens
Selected bacteria's survival rate: 0.566±0.112
Unselected bacteria's survival rate: 0.216±0.096
Interpretations
Interpretations
The data outcomes align with our hypothesis that selection for biofilm formation increases P. fluorescens’ survival rate in response to amikacin injections. Figure 4 shows that the bacteria injected after two rounds of selection survived at a higher rate than those injected without any selection. This difference was statistically significant (p<0.05), and Figure 3, which depicts each survival rate’s colony count breakdown, displays the Day 4 experimental and control groups’ relative proximity compared to Day 2’s, whose control group’s colony count is almost five times greater than its experimental group’s. Although both treatment groups’ difference between control and experimental colony count were statistically insignificant (p>0.05), two of the unselected group’s control replicates were excluded because of overgrowth, and those plates’ absence could account for the unselected group’s statistical insignificance despite the discrepancy between the control and experimental colony counts. Also, Figure 3 was included solely to give context to Figure 4’s survival rates, the experiment’s stated dependent variable. Since the difference in survival rate—which we calculated by considering the colony counts of both the control and experimental groups—between the selected and unselected groups is statistically significant (p~3.239 x 10-5), the results indicate that selection for biofilm formation increases P. fluorescens’ resistance to amikacin.
This interpretation is supported by the roles of biofilms’ extracellular matrix, enzymatic reactions, altered chemical environments, and cells’ stationary phases. When a study tested chlorine’s penetration within a mixed Klebsiella pneumoniae and P. aeruginosa biofilm, the disinfectant’s concentrations in the biofilms were measured at 20% or less of the bulk media’s (1). The chlorine-detecting penetration profile indicated that the weak penetration was a function of chlorine diffusion in the biofilms’ extracellular matrix (1). Likewise, chlorine and other antimicrobial substrates (such as amikacin) are inhibited from killing the bacteria within biofilms because extracellular matrices themselves consume antibiotics. However, the limited amount of antibiotic that does diffuse through the matrix remains vulnerable to degradation from enzymatically based reactions made available by the biofilms’ multicellular structure (2). Besides consumption and degradation, biofilms also facilitate chemical environments that support bacterial survival. Aerobic biofilms, which typically make up a biofilm’s top layers, contain microbial cell clusters and interstitial voids that promote oxygen distribution (3, 4). The voids transport oxygen through biofilms and supply ~50% of the oxygen consumed by cells within them, and biofilms’ complex structure boosts the mass oxygen transport rate: biofilms’ exchange surface is double that of structures characterized by planar geometry (4). P. fluorescens is an obligate aerobe, requiring oxygen to metabolize energy and grow, so living within biofilms structured to distribute sufficient oxygen is crucial to their survival (5). Microbial survival depends on environmental pH as well. When acidity spikes or plummets, bacterial proteins and enzymes can denature, losing their metabolic capacity (6). Single-cell pH analysis has demonstrated that antibiotics increase bacteria’s cytosolic pH, and P. fluorescens are acidophiles that thrive in low pH environments (7). Biofilms mitigate the basic environment that antibiotics induce, though, as the heterotrophs within biofilms compete with nitrifiers for substrate and resist dramatic pH changes (8). This neutralizing effect rescues P. fluorescens from antibiotics that disrupt optimal pH levels for survival. Lastly, biofilms provide a stable environment in which cells can adopt a stationary phase, a slow or non-growth phase of bacteria’s life cycle, which decreases their susceptibility to antibiotic stress (2). Each of these mechanisms—extracellular matrices’ substrate consumption, enzymatic reaction degradation, chemical environment oxidation and pH balancing, and cells’ stationary phases—corroborate biofilms’ role in increasing P. fluorescens’ survival rate in response to antibiotics such as amikacin.
Overall, the data indicate that in response to the antibiotic amikacin, P. fluorescens selected for optimal biofilm formation survive more often than those unselected for optimal biofilm formation, and we propose that this phenomenon occurs because biofilms heighten bacterial antibiotic resistance through four mechanisms: extracellular matrix substrate consumption, enzymatic reactions, balanced chemical environments, and cells’ stationary phases.
Works Cited
Works Cited
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