Cade Burnside, Sam Capehart – Renaissance School, Charlottesville, Virginia, United States of America

Reviewed on 4 May 2024; Accepted on 10 June 2024; Published on 26 October 2024

With help from the 2024 BioTreks Production Team.

Background 

Antibiotic resistance is one of the most pressing medical issues facing modern society. With research projecting that antibiotic resistance will cause 10 million deaths a year by 2050, it is imperative that the scientific community develops new antimicrobials to combat infections (D. M. Lin et al., 2017). One potential solution to the antibiotic resistance crisis is bacteriophage therapy, which uses viruses called bacteriophages to treat bacterial infections. Bacteriophages are the natural predators of bacteria and have had great evolutionary success—they currently number in the tens of nonillions (Zimmer, 2016). The process by which a phage kills its host is relatively simple- the phage first injects its DNA into its target bacterial host, turning the bacterial cell into a phage factory that produces hundreds of phage progenies (Sulakvelidze et. al., 2001). Eventually, the bacterial host cell is lysed, releasing the newly-formed phages out into the environment to continue the infection and propagation cycles. A sufficient body of researchevidence exists in favor of bacteriophage therapy for medical purposes; however, clinical trials using bacteriophages are few and far between, in part because of a severe stigma faced by bacteriophage therapy due to its association with the medical practices of the former Soviet Union (Ireland, 2023). 

Bacteriophages are cheap and easy to produce, store, ship, and administer (Ireland, 2023). They cause few side effects and have been shown to boost the human immune system (Roach et al., 2017). Unlike antibiotics, bacteriophages are host specific. They can be used to only target pathogenic disease-causing bacteria, leaving the helpful bacteria in our microbiome unharmed (Dąbrowska & Abedon, 2019). Furthermore, different strains of bacteriophages are synergistic with one another, with antibiotics, and our immune systems (Segall et al., 2019). They increase immunoglobulin A production in patients and force bacteria to evolve in such a way that the bacteria become more susceptible to antibiotics and other phages (Kuchment, 2010); (Strathdee, 2019).

One of the major concerns about bacteriophage therapy, however, is that bacterial host will evolve and eventually develop resistance to bacteriophages, rendering bacteriophage therapy obsolete in possibly a matter of decades (Dąbrowska & Abedon, 2019). 

Bacterial immunity against bacteriophages can be either innate or adaptive (Hasan & Ahn, 2022). Innate immune responses, which are non-phage-specific, include inhibiting phages from attaching to the cell wall of the bacterial cell, thus preventing the entry of phage DNA into the bacterial cell, weaponizing endonucleases against invading phage DNA, and triggering an “abortive infection system” that sacrifices the infected bacterial cell. Adaptive bacterial immunity, which is phage specific, is initiated by clustered regularly interspaced short palindromic repeats (called CRISPR) and CRISPR-associated proteins (most commonly Cas9). 

The CRISPR/Cas system effectively defends bacteria against bacteriophages by inserting phage-derived DNA sequencesCRISPR into the bacterial host chromosome (Landsberger et al., 2018). When the bacterial cell is invaded by phage DNA, it uses the library of phage DNA stored within its host chromosome to recognize it as a foreign and harmful entity. A guide RNA molecule will then be produced that matches the CRISPR sequence anneals to the phage DNA and directs the endonuclease Cas9 to cleave the phage DNA, preventing it from integrating into the bacterial genome(Anders & Jinek, 2014). 

Phages can counter-evolve to evade CRISPR in a number of ways- point mutations that prevent the bacterial guide RNA from effectively recognizing target phage DNA, anti-CRISPR proteins that inhibit the performance of Cas9, and recombinases that repair the cleaved phage DNA (Hasan & Ahn, 2022). Phages that encode anti-CRISPR (acr) genes can sometimes work together to overcome bacterial immunity, with one phage disabling the host CRISPR-Cas system, while the other phage begins replication within the unfortunate host (Landsberger et al., 2018). There is more research to be done about the exact mechanisms by which bacteriophages can overcome CRISPR/Cas systems, but the ability of phages to rapidly evolve in response to bacterial immunity makes them a more superior and adaptive anti-bacterial treatment solution over antibiotics for treatingfor bacterial infections. 

This experiment aimed to provide insights into what the resistance landscape would look like if bacteriophage therapy was utilized to treat bacterial infections.   Our study consisted of four parts used to compare the ease with which bacteria evolve resistance to antibiotics versus bacteriophages. The first three sections of the experiment determined the optimal conditions for the main part of the experiment, which included finding a growth curve for Escherichia coli (E. coli) DH5α when grown at 37 °C with no agitation, determining the optimal amount of UV exposure to cause maximum mutation in E. coli DH5α, and titrating bacteriophage and antibiotics kanamycin to determine comparable concentrations of these two antimicrobials. Finally, the fourth section compared the growth of two groups of E. coli, which were exposed to UV light and treated with either kanamycin or bacteriophages in order to determine the relative ease at which E. coli evolves resistance to these antimicrobials. The hypothesis was that E. coli would evolve resistance to kanamycin more easily than it did bacteriophage.

The strain of bacteria chosen for this experiment was E. coli DH5α, which is a common, safe, and cheap laboratory strain of bacteria frequently used for its transformation efficiency (Kostylev et al., 2015). Others have performed UV irradiation experiments usingDH5α, supporting its use in this experiment (K. Lin, 2001; J. Zimring, personal communication, October 6, 2023).

The strain of bacteriophage chosen for this experiment was a mutant T4 coliphage called T4r. T4 and T4r are members of the Myoviridae family of bacteriophages and have a long, rigid, contractile tail (Summers, 2001). T4 is one of the seven E. coli phages (T1-T7) discovered by Max Delbrück in 1944 (Yap & Rossmann, 2014). These were the first bacteriophages to be extensively studied, and remain among the most well-understood phages to this day. Additionally, the genes CCTCC AB 2015375responsible for conferring resistance to T4 bacteriophage in E. coli DH5α have been sequenced, making this phage the ideal candidate for testing the development of resistance (Chen et al., 2018). 

This experiment also uses an antibiotic called kanamycin as a control variable. Kanamycin kills bacteria by interfering with the function of their ribosomes. Kanamycin belongs to a family of antibiotics called aminoglycosides and is most frequently used to treat severe tuberculosis infections (PubChem, 2021). Resistance to kanamycin, caused by the pK18mobsacB and pK19mobsacB plasmids, has been observed in bacterial strains such as Mycobacterium tuberculosis, E. coli, and Staphylococcus aureus, among others (Suzuki et al., 1998).

This experiment uses two methods of measuring bacterial growth- standard plate count method and turbidimetric analysis. The standard plate count method was used to measure the number of surviving colonies post-UV radiation and subsequently upon the two treatment conditions. Turbidimetric analysis was used to develop the growth curve for E. coli and measure growth when the bacteria were exposed to either kanamycin and bacteriophage. 

Turbidimetric analysis is performed using a machine called a spectrophotometer, which can be used to measure the incident light scattered by bacteria in a liquid culture. The resulting optical density is proportional to the number of bacteria in the solution. Cultures of E. coli are traditionally measured using light with a wavelength of 600 nanometers (abbreviated A₆₀₀) (Du et al., 2022).

The optical density of a bacterial culture is used to determine what phase of growth the bacteria are in. Bacteria grow exponentially, and the life cycle of a bacterial culture can be divided into four phases- lag, log, stationary, and death (Maier, 2009). The initial slow-growing lag phase is followed by the log phase, which is characterized by an extreme increase in population (Bruslind, 2018). In the stationary phase, growth slows and gradually evens out until the death phase, when all of the available nutrients have been consumed and the bacteria population starts to die. Bacterial cells are the healthiest and most uniform during the late log phase, so the majority of experiments are performed during this phase. This particular experiment was performed when the bacterial culture reached an optical density of 0.7 at A₆₀₀, following established standards for ultraviolet mutagenesis (K. Lin, 2001).

Ultraviolet (UV) light is light with a wavelength between 100 and 400 nanometers (CDC, 2023). It is an established means of disinfection, frequently used to sterilize the air in hospitals, restaurants, and schools (Reed, 2010). UV light kills bacteria by interfering with their DNA (Pereira et al., 2014). When a bacterial cell absorbs an ultraviolet photon, it can cause erroneous bonds to form between neighboring nitrogenous bases of DNA, mutating genes and eventually rendering the bacteria incapable of replicating. In the attemptto fix these UV-induced lesions, the error-prone DNA polymerase enzyme PolV of the bacterial introduces several random base pairs into the DNA strand opposite the lesion (Krishna et al., 2007). This causes mutations in the bacterial genetic code, allowing for E. coli to evolve and develop resistance (among other things) more quickly.

This experiment used a Black Magic 25 W germicidal UV lamp. An ozone-free lamp was chosen because ozone can also interfere with bacterial growth (Rangel et al., 2021). This experiment sought to determine the optimal UV exposure time for creating significant mutations without increased bacterial death. The optimal time was considered to be the longest exposure time that still retained a “bacterial lawn” —more than 300 living colonies inhabiting the same plate.

Materials and methods

Materials

Agar plates, bunsen burner, cuvettes, E. coli DH5α (Carolina Biological Supply), hot plate, 35 mg/mL kanamycin (Carolina Biological Supply), Luria Broth (LB), micropipettes, microplate, pipette tips, spectrophotometer, T4r bacteriophage (Carolina Biological Supply), test tubes, tinfoil, ultraviolet lamp (Black Magic) and water.

Growth curve   

An overnight culture of E. coli DH5α was grown in a ratio of 1:200 (10 µl E. coli suspension to 2 mL Luria Broth). The next day, 5 mL of LB was inoculated with 100 µl of the overnight culture and placed in a 37°C incubator to grow without agitation. Every hour, the culture was removed from the incubator and shaken by hand. Three mL of the E. coli culture were pipetted into a sterile cuvette. The spectrophotometer was blanked with a cuvette full of LB, and the optical density of the E. coli was measured at wavelengths of A₆₀₀ and A₆₆₀, before the culture was pipetted back into the test tube and returned to the incubator for another hour. This procedure was repeated a total of ten times to form a growth curve for the log phase of E. coli DH5α under these conditions (Fig. 2).

Figure 2. A graph showing the growth curve of E. coli DH5α at a wavelength of A600.

UV irradiation

E. coli from an overnight culture was grown in LB in a 1:50 ratio until it reached an optical density of 0.7 at A₆₀₀. Eight agar plates were labeled 1-8, and 100 µl of E. coli was spread-plated onto plates 1-7. Irradiation of the plates was performed with a 25W ozone-free UV lamp at a vertical distance of 28cm, following a similar experiment described by Hong et al., 2016. The lamp was warmed up for 30 minutes prior to irradiation (K. Lin, 2001). Plates were then placed beneath the lamp for a predetermined duration (15s, 30s, 45s, 60s, 75s, 90s, 105s, 120s, 0s) with the lids of the plates removed to avoid any shielding effect. Plate 8, which had no E. coli on it, was exposed to the UV lamp for the entire duration of the experiment to ensure that nothing contaminated the other plates while they were being irradiated (negative control). After irradiation, the plates were incubated in a 37°C incubator for 24 hours. The next day, the plates were photographed and the number of E. coli colonies present was counted and recorded (Fig. 3).

Figure 3. A graph showing the number of colonies present in plates 1-7 after 24 hours of incubation.

Phage titration

The first five wells of a sterile microplate were each filled with 2 mL of LB. Well 1 was inoculated with 200 µl of T4r bacteriophage and pipetted up and down several times to mix. One hundred microliters of solution was transferred from well 1 into well 2, and the solution was mixed. This step was repeated for wells 3 and 4 to form a two-fold dilution. One hundred microliters of solution was discarded from well 4, and well 5 was left as the negative control (Fig. 1). Five milliliters of E. coli DH5α overnight culture was grown in LB in a 1:50 ratio until it reached an optical density of 0.7 at A₆₀₀. Ten microliters of E. coli were then pipetted into each well. 

The microplate was placed in a 37° C incubator and left to incubate overnight. The next day, the optical densities of wells 1-5 were measured in a spectrophotometer at A₆₀₀ (see Figure 4).

Figure 1. An image showing the two-fold dilution series in wells 1-5 of the phage titration.

Resistance comparison

Five milliliters of E. coli DH5α was grown in LB in a 1:50 ratio until it reached an optical density of 0.7 at A₆₀₀. The first ten wells of two sterile microplates were each filled with 2 mL of LB. The first nine wells of both microplates were each inoculated with 10 µl E. coli. One microplate was designated the control plate, the other was irradiated by a 25W UV lamp at a 28cm distance for 90 seconds. The lamp was allowed to warm up for 30 minutes prior to irradiation. 

Following irradiation, both microplates were inoculated with either kanamycin orbacteriophage. The ratio of kanamycin/bacteriophage to bacteria were as follows:

Kanamycin wells:

K1: 1 µl kanamycin (35 mg/mL), 10 µl E. coli

K2: 0.5 µl kanamycin, 10 µl E. coli 

K3: 0.25 µl kanamycin, 10 µl E. coli

K4: 0.1 µl kanamycin, 10 µl E. coli

Bacteriophage wells:

P1: 200 µl bacteriophage, 10 µl E. coli

P2: 150 µl bacteriophage, 10 µl E. coli

P3: 50 µl bacteriophage, 10 µl E. coli
P4: 50 µl bacteriophage, 10 µl E. coli

The ninth well was not inoculated with an antimicrobial (untreated control). The tenth was not inoculated with anything (no bacteria, no treatment control). Both the ninth and tenth wells served as negative controls. Both microplates were left in a 37°C incubator for 24 hours. The next day, solutions were diluted and their optical densities were measured in a spectrophotometer at A₆₀₀ (Fig. 5). The difference between irradiated and non-irradiated cultures was calculated (Fig. 6). The data from each individual concentration was placed into a table to be analyzed (Table 1, Table 2).  This procedure was repeated twice (see Fig. 7, Fig. 8, Table 3, Table 4). Extensive measures were undertaken to ensure the validity of the experiment. Sterile equipment was used at every stage to eliminatethe risk of contamination. Every section of the experiment was repeated a minimum of three times, and positive and negative controls were present at each step. The experiment was performed on 25 individual days across a two-month period. 

Results

 E. coli DH5a reaches 0.7 OD at ten hours

The growth curve determined that a 50:1 ratio of E. coli DH5α to LB, grown at 37°C without agitation in a 20 mL test tube reached the desired optical density of 0.7 at A₆₀₀ after 10 hours of growth (Fig. 2). This determined that E. coli would be grown for 10 hours during subsequent sections of the experiment, because 0.7OD is the optimal optical density for E. coli mutagenic irradiation with UV light. 

Figure 2. A graph showing the growth curve of E. coli DH5α at a wavelength of A600.

 The optimal UV irradiation time for E. coli mutagenesis is 90s

Six separate irradiation experiments indicated that 90 seconds was the optimal irradiation time. All of the plates exposed to fewer than 90 seconds of UV light showed relatively high numbers of colonies, but the colony count dropped drastically after 90 seconds of exposure (Fig. 3). This determined that 90 seconds of irradiation time would be used in the experiment moving forward, because this time allowed for maximum bacterial mutagenesis without killing large amounts of the bacterial culture.

Figure 3. A graph showing the number of colonies present in plates 1-7 after 24 hours of incubation.

Phage Titration Results

 A turbidimetric analysis of the liquid cultures from the initial phage titration showed an optical density of E. coli ranging from 0.63 (a  20:1 ratio of phage to bacteria) to 0.96 (the negative control, which was not inoculated with any phage) (Fig. 4). The initial phage dilution was two-fold, but the resulting optical density of the E. coli was not exactly halved —this is likely because bacteriophages are self-propagating. Once a certain saturation of bacteriophage was reached, the addition of more bacteriophages became redundant. This phage titration determined that a 0.1:1 ratio of kanamycin to E. coli, as determined by Garza-Cervantes, 2020, killed roughly as much bacteria as a 20:1 ratio of bacteriophage to E. coli did.  This rough equivalency was used to determine the titrations of bacteriophage and kanamycin for the fourth and final section of the experiment.

Figure 4. A graph showing the initial bacteriophage titration. The optical density of the E. coli is on the y-axis, and the ratio of bacteriophage to E. coli is on the x-axis.

 Following irradiation, E. coli resists kanamycin more successfully than bacteriophage

We observed that kanamycin killed more non-irradiated E. coli than irradiated E. coli, whereas bacteriophages were slightly more successful at killing the irradiated version of the bacteria. While this cannot be extrapolated to a clinical setting, our results showed that bacteriophage therapy has the potential to be more effective in treating bacterial infections as bacteria do not develop resistance to phages as quickly as to antibiotics.  In the first resistance comparison, 75% of the irradiated kanamycin-treated cultures grew more than the control. The growth of kanamycin-treated E. coli increased by an average optical density of 0.19 in the irradiated group when compared to control. All of the irradiated phage-treated cultures grew less than the control group of phage-treated cultures. The growth of phage-treated E. coli decreased by an average optical density of 0.16 when compared to the control.

Figure 5. Trial one- A graph showing the optical density of E. coli DH5α when exposed to different concentrations of kanamycin (K1-K4) or bacteriophage (P1 – P2) and either irradiated (red) or not irradiated (blue). The two columns on the far right, labeled C, represent the control group.

Figure 6. A graph showing the difference in E. coli growth between the control group and the irradiated group at each concentration of kanamycin or bacteriophage.

Table 1. Trial one- A table showing the optical density of the control group and irradiated group for each concentration of kanamycin, as well as the difference between the two groups and the average value.

Ratio of kanamycin to E. coli	OD of Control Group	OD of Irradiated Group	Difference (Irradiated – Control)
0.1:1	0.71	1.1	0.39
0.05:1	1.0	1.44	0.44
0.025:1	1.17	0.93	-0.24
0.01:1	1.1	1.26	0.16
Average:	1.0	1.18	0.19

Table 2. Trial one- A table showing the optical density of the control group and irradiated group for each concentration of bacteriophage, as well as the difference between the two groups and the average value.

Ratio of phage to E. coli	OD of Control Group	OD of Irradiated Group	Difference (Irradiated – Control)
20:1	1.45	1.0	-0.45
15:1	1.03	0.99	-0.04
10:1	1.0	0.93	-0.07
5:1	1.08	0.99	-0.09
Average:	1.14	0.98	-0.16

In the second trial, we observed consistent results. The growth of kanamycin-treated E. coli once again increased by an average optical density of 0.19 (Table 3), with 75% of irradiated cultures growing more than their non-irradiated counterparts (Fig. 7, 8). The growth of bacteriophage-treated E. coli decreased by an average optical density of 0.05 ( Table 4), with a 50-50 split between irradiated cultures that grew more or less than their counterparts (Fig.7, 8). 

Table 3. Trial Two: A table showing the optical density of the control group and irradiated group for each concentration of kanamycin, as well as the difference between the two groups and the average value.

Ratio of kanamycin to E. coli	OD of Control Group	OD of Irradiated Group	Difference (Irradiated – Control)
0.1:1	0.92	1.17	0.25
0.05:1	1.11	1.44	0.33
0.025:1	0.94	0.9	-0.04
0.01:1	0.95	1.19	0.24
Average:	0.98	1.18	0.19

Figure 7. Trial two- A graph showing the optical density of E. coli DH5α when exposed to different concentrations of kanamycin (K1-K4) or bacteriophage (P1 – P2) and either irradiated (red) or not irradiated (blue). The two columns on the far right, labeled C, represent the control group.

Figure 8. Trial two- A graph showing the difference in E. coli growth between the control group and the irradiated group at each concentration of kanamycin or bacteriophage.

Table 4. Trial two- A table showing the optical density of the control group and irradiated group for each concentration of bacteriophage, as well as the difference between the two groups and the average value.

Ratio of phage to E. coli	OD of Control Group	OD of Irradiated Group	Difference (Irradiated – Control)
20:1	1.24	1.32	0.08
15:1	1.2	1.03	-0.17
10:1	0.9	0.93	0.03
5:1	1.1	0.98	-0.12
Average:	1.11	1.07	-0.05

The increase of growth in kanamycin-treated cultures indicates that the UV light mutated the E. coli genome in such a way that the bacterial culture became genetically diversified and began to evolve resistance to the kanamycin. Although both the kanamycin-treated and phage-treated E. coli cultures were exposed to the same duration of UV light, the irradiated phage-treated cultures did not outperform the non-irradiated phage-treated cultures. This indicates that the bacteriophages were able to evolve at a similar rate to the irradiated E. coli, preventing the bacteria’s UV-induced genetic diversity from giving it an evolutionary advantage. On the other hand, the UV-induced mutagenesis of the bacterial genome allowed it to develop resistance against kanamycin more effectively. 

Figure 9. A table showing the calculations performed in order to determine the t-score and p-value for the experiment.

The change in optical density from irradiated samples compared to controls was calculated for each antimicrobial concentration in both trials. A t-test was performed on this data in order to determine whether the difference between the kanamycin group and the phage group was statistically significant: t(7) = 2.98, p = 0.0099. The p-value of 0.0099 gives a >99.99% confidence that this data was not generated due to random chance. This supports our hypothesis that E. coli DH5α will not evolve resistance to bacteriophages as easily as it does kanamycin, because the bacteriophages have the ability to evolve in parallel.

Discussions

Herein, we have shown that the  results of this experiment are promising, but  there is much more to be explored in the field of bacterial resistance to bacteriophages, including research into innate bacterial immune responses, the CRISPR/Cas system, and how phages combat bacterial defense mechanisms. 

It would be ideal to expound upon the results of this experiment in greater detail by determining how E. coli and phage interact on the molecular level following UV light exposure. The experiment would also benefit from being repeated with a larger sample size, more sensitive equipment, and multiplestrains of bacteria, and phage, as well as other types of antibiotics. If the results stay consistent, this experiment could eventually be adapted to help make advancements in the understanding of phage-antibiotic synergy.  

Next steps  

When contemplating the introduction of bacteriophage therapy into mainstream Western medicine, the threat of bacterial resistance to bacteriophages is not something to be overlooked. However, our results support the idea that bacterial resistance to bacteriophages should not be a factor to completely dismiss the therapeutic potential of bacteriophage therapy in combating bacterial infections  This experiment showed that there is a statistically significant difference in the ease with which E.coli DH5α evolves resistance to kanamycin when compared to the ease with which it evolves resistance to T4r bacteriophage. Following UV-induced mutagenesis, phage-treated E. coli grew significantly less than kanamycin-treated E. coli. This indicates that the ability of bacteriophages to evolve alongside their hosts may help mitigate the development of bacteriophage resistance. Furthermore, the natural evolution of phages is only one of the many ways to decrease risk of developing resistance;  researchers can look into improving phage activity by genetically engineering new phages, creating synergistic cocktails, using phages in tandem with antibiotics, and isolating lytic enzymes produced by phages to develop new therapies. 

Bacteriophage therapy has a notorious history However, as more and more experiments like ours demonstrate the vast potential of bacteriophages, the scientific community will hopefully begin to recognize bacteriophage therapy for what it is: the singular most promising solution to the ongoing antibiotic resistance crisis. 

Author contributions

SFC performed bulk of the experimentsn, while CSB performed tasks such as pouring up agar plates and inoculating liquid media. SFC wrote the original draft of the paper and created the graphs and tables. CSB reformatted the paper and aid in revising the graphs and tables.

Acknowledgments

The authors would like to acknowledge Ms. Anna Minutella and the Renaissance High School, who provided experimental design critique, materials, and a laboratory space in which to perform the experiment. The authors would also like to acknowledge Dr. James Zimring from the University of Virginia, who contributed his knowledge about pathology, spectroscopy, and experimental design, and assisted in the selection of E. coli DH5α, T4r bacteriophage, and kanamycin as the experimental materials. The authors would also like to thank Ollie Munzenrider, another Renaissance student who helped us start this project.

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