We thank Gor Margaryan for creative support and technical assistance. This work was supported by the ZMB Young Scientist Grant 2017/2018 (awarded to TW) and the DFG focus program 1819 (Grant No. DA1202/2-1 awarded to TD).

1. Construction of a model plasmid carrying an antibiotic resistance gene
NOTE: The strain Escherichia coli K-12 MG1655 was used as the model organism in all experiments (DSM No. 18039, German Collection of Microorganisms and Cell Cultures, DSMZ). The strain E. coli DH5&#945;17 was used during plasmid construction.
<ol>
	<li>PCR amplify the plasmid backbone of your choice and amplify the resistance gene including the promoter region by PCR (Figure 1).
	<ol>
		<li>PCR amplify the plasmid backbone using a high-fidelity polymerase and the oligonucleotides pBBR1_for (5&#39;-GCGGCCACCGGCTGGCT-3&#39;) and pBBR1_rev (5&#39;-TACCGGCGCGGCAGCGTGACCC-3&#39;) on the plasmid template pLC (GenBank acc. no. MH238456)18.</li>
		<li>PCR amplify the resistance gene nptII including the native Tn5 promoter19 using a high-fidelity polymerase and oligonucleotides nptII_gib_for (5&#39;-GCGCCGGTAGATCTGCTCATGTTTGAAGCTTCACGCTGCCGCA-3&#39;) and nptII_gib_rev (5&#39;-CGGTGGCCGCCAAAAAGGCCATCCGTCAGGTCAGAAGAACTCGT-3&#39;). The nptII gene encodes for a neomycin phosphotransferase and confers resistance to kanamycin. 
		NOTE: Design the primers for the resistance gene with approximately 20 bp of complementary sequence to the plasmid backbone that it will be fused to.</li>
	</ol>
	</li>
	<li>Clean both PCR fragments using a kit of your choice.</li>
	<li>Join the purified antibiotic resistance gene PCR product (including its promoter region) to the purified plasmid backbone and fuse the homologous regions using isothermal assembly20, at 50 &#176;C for 60 min.</li>
	<li>Electroporate the fused product into the strain E. coli DH5&#945;.
	<ol>
		<li>Introduce 2 &#181;L of the product from step 1.3 into 40 &#181;L of electrocompetent cells in 2 mm cuvettes at 4 &#176;C and 2.5 kV. Resuspend cells in 1 mL of lysogeny broth (LB) medium.</li>
		<li>Transfer the total volume to a microfuge tube and incubate 1 h at 37 &#176;C shaking at 250 rpm in an orbital shaker to allow for the expression of the resistance marker on the plasmid.</li>
		<li>Plate 100 &#181;L of the cells on LB agar plates containing the appropriate antibiotic (kanamycin 25 &#181;g/mL) to select for the antibiotic resistance gene and thus select for plasmid-carrying cells. Spin down the rest, remove the supernatant, resuspend the cells in 100 &#181;L LB and plate on a selective agar plate. Incubate the plates at 37 &#176;C for 24 h.</li>
	</ol>
	</li>
	<li>In order to verify clones, extract the constructed plasmids via alkaline lysis using a commercial mini-prep kit.
	<ol>
		<li>Harvest 5 mL of the stationary overnight culture by centrifuging at 12,000 x g at room temperature. Resuspend the cells in resuspension solution, then lyse and neutralize the cell solution. Centrifuge for 5 min at 12,000 x g.</li>
		<li>Transfer the supernatant to the DNA binding column provided in the kit and wash the column twice with 500 &#181;L washing solution centrifuging 2x before eluting the column membrane with elution buffer.</li>
		<li>Perform Sanger sequencing of the plasmid to confirm that the sequence is correct.</li>
	</ol>
	</li>
	<li>Once the plasmid validity is verified, electroporate the plasmid (now pCON) into the strain E. coli MG1655 as described above. This yields strain E. coli MG1655 pCON.</li>
</ol>
2. Monitoring plasmid-carrying bacteria under various conditions over time
NOTE: The evolution experiment is conducted with plasmid-carrying strains under nonselective conditions (LB media) in two temperatures (37 &#176;C and 20 &#176;C) and three population bottleneck sizes. The experimental design is used to study plasmid persistence under various conditions.
<ol>
	<li>Design of an evolution experiment to follow plasmid frequency over time (Figure 2)
	<ol>
		<li>Plate the constructed plasmid-carrying strain (E. coli MG1655 pCON) on LB agar plates supplemented with antibiotics (kanamycin 25 &#181;g/mL) and incubate overnight at 37 &#176;C.</li>
		<li>Prepare 96 deep-well plates with 1 mL of LB medium in each well. As the bacterial ancestors, pick eight random isolated colonies from the agar plate in independent wells. Incubate the plates at 37 &#176;C and 450 rpm on a plate shaker for 24 h. Prepare a frozen glycerol stock of the ancestral clones.</li>
		<li>The next day transfer the eight replicate populations into new deep-well plates according to the experimental design (Figure 2). The cultures are diluted 1:100 (large bottleneck, L), 1:1,000 (medium bottleneck, M), or 1:10,000 (small bottleneck, S) in a total volume of 1 mL LB using PBS for dilution. The diluted cultures are both incubated at 37 &#176;C and 20 &#176;C. 
		NOTE: It is highly important to control for cross-contamination in the 96-deep-well plate. Thus, use a checkerboard plate design by intercalating inoculated wells with bacteria-free LB medium. Use this pattern through the entire evolution experiment.</li>
		<li>The cultures incubated at 37 &#176;C are transferred every 12 h while cultures at 20 &#176;C are transferred every 24 h. 
		NOTE: During every transfer event the bottleneck size treatment is applied and the serial transfer is repeated over a total of 98 transfers. The number of transfers will depend on the readers&#39; experimental design.</li>
		<li>Prepare a frozen glycerol stock of all the populations regularly, 2x a week.</li>
	</ol>
	</li>
	<li>Monitoring the plasmid frequency by replica plating (Figure 3) 
	NOTE: During the evolution experiment, the frequency of plasmid-carrying cells in the population is estimated from the proportion of hosts. The replica plating protocol is shown in Figure 3.
	<ol>
		<li>To determine the plasmid frequency in the population during the evolution experiment, stationary cultures are serially diluted and plated on nonselective LB agar plates. Adjust the dilution according to a yield of 250-500 colonies per plate. 
		NOTE: Prepare thick LB agar plates prior to the plating (~30 mL agar).</li>
		<li>The plated populations are incubated for overnight growth according to their growth temperature in the experiment. 
		NOTE: The colonies need to be small, thus incubate &#60;24 h at 37 &#176;C.</li>
		<li>After overnight growth, count all colonies using a manual or automated colony count station and calculate the total bacterial population size in the cultures. 
		NOTE: Colonies at the edge of the agar plate should not be included in the total bacterial cell count.</li>
		<li>Sterilize square pieces (~20 x 20 cm) of cotton velvet by autoclaving. 
		NOTE: The velvet cloth needs to be 100% cotton to be autoclavable. It is important to dry the cloth after sterilization.</li>
		<li>After sterilization of the velvet cloth, place the cloth on a round block and fix it with a metal ring. It is crucial to avoid wrinkles. Carefully place the plate with the grown colonies (agar facing down) on the fixed velvet cloth surface. Make sure that all colonies touch the velvet surface by carefully tapping on the Petri dish in a circular manner.</li>
		<li>Carefully remove the LB agar plate and place a selective plate supplemented with antibiotics (kanamycin 25 &#181;g/mL) onto the velvet cloth. Make sure the plate is touching all the velvet by carefully tapping on the Petri dish as described before. Afterwards, remove the plate. Leave the plates at room temperature for overnight growth.</li>
		<li>The next day, evaluate both the LB agar plate and the selective plate. Colonies that grow on the selective media are counted as plasmid hosts (i.e., antibiotic resistant), while colony-free spots are colonies that were plasmid-free and are thus not resistant to the antibiotic (i.e., lost the plasmid). This is done by placing the plates over each other and comparing the growth (i.e., mark any missing colonies) and counting the colony number on both plates. This yields the number of cells that lost the plasmid during the evolution experiment.</li>
		<li>Repeat this procedure along the whole evolution experiment in a regular manner (e.g., every 14 transfers).</li>
	</ol>
	</li>
</ol>
3. Visualization of plasmid multimers using gel electrophoresis
NOTE: Plasmid extraction of low-copy plasmids often leads to contamination with host chromosomal DNA that needs to be enzymatically digested prior to visualization.
<ol>
	<li>Extract plasmid DNA from a 5 mL stationary overnight cell culture using alkaline lysis as described in step 1.5.</li>
	<li>Afterwards, treat the extracted plasmid DNA with an ATP-dependent DNase that only cuts chromosomal DNA to remove chromosomal DNA contamination (see Table of Materials). Incubate at 37 &#176;C for 30 min. Afterwards, clean the DNA using a kit of your choice.</li>
	<li>To create open circle molecules of all plasmid conformations (monomers or multimers), incubate the plasmid DNA samples with a nicking enzyme (Nb.BsrDI) and incubate for 30 min at 65 &#176;C.</li>
	<li>In parallel, to create linear plasmid molecules (i.e., for plasmid size comparison), use a restriction enzyme of your choice (e.g., HindIII) that cleaves the plasmid once. 
	NOTE: This results in linear DNA plasmid monomers. Open circle molecules do not migrate in a linear manner.</li>
	<li>To visualize the plasmid size and conformation, electrophorese the nicked and linear plasmid DNA samples for 120 min at 4.3 V/cm in a 1% (w/v) agarose gel and 1&#215; TAE buffer. The samples are stained with Midori green and the gel is visualized on a gel imaging system (see Table of Materials). Use a 1 kbp ladder.</li>
</ol>

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The authors have nothing to disclose.

In this protocol, we present an approach that combines techniques in molecular biology, experimental evolution, and DNA visualization to investigate the role of plasmid evolution for the persistence of antibiotic resistance in bacteria. Although the presented approach combines methods from different research areas, all the applied techniques are straightforward and can be performed in a standard microbiology laboratory.
The most critical steps in the protocol include the construction of the model system strain that includes the genetic validation of the plasmid-carrying genotype. Notably, many plasmids naturally encode antibiotic resistance genes. Thus, the reader may omit step 1 of the protocol and directly proceed with step 2. Next, the evolution experiments should include a random design of replicate populations so that the results are not biased by the position of the replicate populations in the deep-well plate. In addition, it is especially important to carefully conduct the serial transfer and dilution steps in the evolution experiment as contamination would falsify the results. Lastly, replica plating should be conducted with great care. Large colony size may be an issue, but it can be avoided by incubating the plates for less than 24 h. Similarly, the number of colonies on one plate might bias replica plating results. Therefore, populations need to be diluted prior to plating and replication.
One of the biggest advantages of our approach is that is can be easily reproduced without the need for heavy equipment. In addition, another advantage of replica plating to follow marker genes is that only live cells are evaluated, in contrast to flow cytometry or qPCR in which dead cells may be evaluated as alive. Thus, replica plating introduces less bias to the counting of plasmid-carrying cells. Nonetheless, one limitation of replica plating may be the population size (i.e., cell number) that is possible to evaluate in one experimental run.
Using our approach, we have recently shown that plasmid stability evolution potentiates the persistence of antibiotic resistance genes in bacteria. Thus, we developed an approach as a tool to follow plasmid-mediated resistance persistence that is of high importance to follow resistance over time especially under conditions without the presence of antibiotics.

Plasmids are circular, self-replicating genetic elements that are ubiquitous in prokaryotes. They are agents of lateral gene transfer, as they can transfer traits between microbial populations, and thus are considered to play a major role in microbial evolution. Plasmids are drivers of rapid adaptation to growth-limiting conditions over a short time (e.g., in the presence of antibiotics or pesticides1) and are responsible for long-term transition to other lifestyle modes (e.g., emergence of pathogenicity2). The most striking examples for the impact of plasmids on transfer of genes are documented in ecosystems exposed to fluctuating levels of antibiotics, such as medical clinics or in industrial farms3. Due to strong positive selection, many plasmids encode for antimicrobial resistance genes and are often found to confer multiresistance to their bacterial host. Plasmids enable migration between populations or bacterial species, resulting in a rapid propagation of multiple antimicrobial resistance. Under nonselective conditions plasmids are not essential to the cell and are often even referred to as parasitic elements. Nonetheless, plasmids are ubiquitous in nature and their evolution is highly intertwined with that of bacterial chromosomes. Plasmid persistence in natural environments (fluctuating and nonselective) remains poorly understood, yet it is of high importance for our understanding of the persistence of antibiotic resistance genes in nature.
Experimental evolution is a powerful tool for the study of microbial populations4. Experimental evolution demonstrated that imposing strong selection for plasmid maintenance leads to compensatory (i.e., adaptive) evolution of the plasmid or host chromosome that reduces the plasmid fitness cost and, in turn, facilitates plasmid abundance (i.e., plasmid persistence)5,6,7. Thus, following the plasmid-host interaction over time may reveal important mechanisms of adaptation of both elements. Furthermore, experimental evolution enables one to quantify the abundance of plasmid-carrying cells over time under various conditions8,9,10.
Plasmid persistence in evolution experiments can be monitored by several strategies including flow cytometry by fluorescent activated cell sorting (FACS)11, quantitative PCR (qPCR)11, or in cultivation-based methods. Flow cytometry requires a FACS machine and the introduction of a detectable (fluorescent) marker gene, such as the green fluorescent protein (GFP), on the plasmid. However, GFP expression may alter several cellular properties and furthermore influence the plasmid location in the cell12, which in turn may influence plasmid inheritance during cell division. A qPCR approach to measure plasmid abundance may be highly biased by the plasmid copy number, which can vary greatly along bacterial growth phase and over time13. Lastly, a culture-based and plating approach requires the introduction of a selectable marker gene. This may be an antibiotic resistance gene, which is often encoded on natural plasmids; thus, no genetic manipulation is necessary. Antibiotic resistance may be followed by a traditional replica plating approach. Thus, to study natural plasmid dynamics, replica plating is well-suited to monitor plasmid-encoded antibiotic resisitance14.
To visualize plasmid molecules (e.g., to evaluate plasmid size) several methods may be applied. Whole plasmids can be amplified using a PCR-based approach. However, this requires the design of specific primers, which may be challenging during an evolution experiment, because the plasmid sequence may change over time. In addition, it is difficult to amplify plasmid multimers in a PCR-based approach due to multiple binding sites for the PCR primers. Multimeric plasmid molecules can appear following plasmid replication termination or through recombination of plasmid molecules and are mostly oriented head-to-tail15. Another approach of plasmid visualization combines enzymatic digestion of plasmid molecules by DNA endonucleases that either cleave or nick a plasmid DNA strand with agarose gel electrophoresis analysis. The same plasmid of different sizes (e.g., monomers vs. multimers) results in different gel mobilities that can be observed when visualizing the plasmid molecules. This approach enables visualization and quantification of different plasmid conformations (i.e., multimerization states). The plasmid conformation may be used as an indicator of plasmid stability, as plasmid multimers are frequently lost during cell division16.
In a recent work, we followed plasmid persistence in conditions that were not selective for plasmid abundance (i.e., without antibiotic selection). We compared plasmid persistence at two different temperatures (20 &#176;C and 37 &#176;C) and three population sizes (i.e., dilution rates). Applying various dilution rates, or population bottlenecks, allows for the investigation of the influence of population size on bacterial and plasmid evolution. Based on our results, we propose that plasmids can be neutral to their bacterial host and may evolve stability without any selection pressure8. The evolved plasmid stability is conferred by the reduction of plasmid multimer formation8.
Here, we present a protocol for the quantification of plasmid persistence and investigation of plasmid evolution in regard to the maintenance of antibiotic resistance genes. The method has several steps, including the insertion of an antibiotic resistance gene to a model plasmid (which can be omitted when using a natural resistance plasmid), followed by the use of experimental evolution to assess the potential of the plasmid to persist under nonselective conditions while determining the plasmid frequency dynamics over time using replica plating, and the analysis of the plasmid genome by visualization. The protocol described here was designed to investigate the evolution and persistence of plasmids, but it may also be applied to follow the evolution of chromosomal resistance genes (or other marker genes) over time.

<table><tbody><tr><td>96-deep-well plates</td><td>Starlab</td><td></td><td></td></tr><tr><td>96-deep-well plates</td><td>Roth</td><td>EN07.1</td><td>2 ml, square</td></tr><tr><td>96-deep-well plates (cryo)</td><td>Starlab</td><td>E1702-8400</td><td>Micro-Dilution Tube System</td></tr><tr><td>Colony counter</td><td>Stuart</td><td>SC6+</td><td></td></tr><tr><td>Cotton velvet</td><td>drapery shop</td><td></td><td>100 % cotton required</td></tr><tr><td>Electrophoresis chamber</td><td>BioRad</td><td></td><td>Agarose gel electrophoresis</td></tr><tr><td>Electrophoresis power supply</td><td>BioRad</td><td>1645070</td><td>Agarose gel electrophoresis</td></tr><tr><td>Electroporation cuvettes</td><td>BioRad</td><td>1652089</td><td>0.1 cm</td></tr><tr><td>Electroporator</td><td>BioRad</td><td>1652660</td><td></td></tr><tr><td>GeneJet Gel Extraction kit</td><td>Thermo Fisher Scientific</td><td>K0832</td><td>PCR fragment clean-up</td></tr><tr><td>GeneJet Plasmid Miniprep kit</td><td>Thermo Fisher Scientific</td><td>K0503</td><td>Plasmid extraction kit</td></tr><tr><td>Gibson Assembly</td><td>New England Biolabs</td><td>E2611S</td><td></td></tr><tr><td>Incubator</td><td>Thermo Fisher Scientific</td><td>50125852</td><td></td></tr><tr><td>Incubator (plate shaker)</td><td>Heidolph</td><td>1000</td><td></td></tr><tr><td>Incubator (shaker)</td><td>New Brunswick Scientific</td><td>Innova 44</td><td></td></tr><tr><td>Inoculating loops</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Multi-channel pippetes</td><td>Eppendorf</td><td>3125000052, 3125000028</td><td></td></tr><tr><td>Multi-channel pippetes</td><td>Capp</td><td>ME8-1250R</td><td></td></tr><tr><td>NanoDrop 2000/2000c</td><td>Thermo Fisher Scientific</td><td>ND2000</td><td></td></tr><tr><td>Oligonucleotides</td><td>Eurofines</td><td></td><td></td></tr><tr><td>Petri dishes</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Phusion Polymerase</td><td>Thermo Fisher Scientific</td><td>F533S</td><td></td></tr><tr><td>Pipettes</td><td>Eppendorf</td><td>3123000012, 3123000098, 3123000055, 3123000063,</td><td></td></tr><tr><td>PlasmidSafe enzyme</td><td>Epicentre</td><td>10059400</td><td></td></tr><tr><td>Reaction tubes</td><td>Eppendorf</td><td>30125150</td><td></td></tr><tr><td>Replica block &amp;amp; metal ring</td><td>VWR</td><td>601-3401</td><td>PVC cylinder 69 mm; ring 102cm</td></tr><tr><td>Resctriction enzymes</td><td>New England Biolabs</td><td></td><td></td></tr><tr><td>Thermocycler</td><td>BioRad</td><td>T100</td><td></td></tr></tbody></table>

quantification of plasmid-mediated antibiotic resistance in an experimental evolution approach

Plasmids play a major role in microbial ecology and evolution as vehicles of lateral gene transfer and reservoirs of accessory gene functions in microbial populations. This is especially the case under rapidly changing environments such as fluctuating antibiotics exposure. We recently showed that plasmids maintain antibiotic resistance genes in Escherichia coli without positive selection for the plasmid presence. Here we describe an experimental system that allows following both the plasmid genotype and phenotype in long-term evolution experiments. We use molecular techniques to design a model plasmid that is subsequently introduced to an experimental evolution batch system approach in an E. coli host. We follow the plasmid frequency over time by applying replica plating of the E. coli populations while quantifying the antibiotic resistance persistence. In addition, we monitor the conformation of plasmids in host cells by analyzing the extent of plasmid multimer formation by plasmid nicking and agarose gel electrophoresis. Such an approach allows us to visualize not only the genome size of evolving plasmids but also their topological conformation-a factor highly important for plasmid inheritance. Our system combines molecular strategies with traditional microbiology approaches and provides a set-up to follow plasmids in bacterial populations over a long time. The presented approach can be applied to study a wide range of mobile genetic elements in the future.

Here, we present an approach to study plasmid evolution by quantifying plasmid persistence in a population. First, we show how to build the plasmid carrying E. coli strain MG1655 pCON that is subsequently introduced to an evolution experiment. Second, we present a straightforward method to follow plasmid abundance in the evolving bacterial populations. Finally, we show how to visualize plasmid molecule size and conformation.
In our previous work8 using the presented approach we conducted an evolution experiment following the persistence of antibiotic resistance plasmids in E. coli in the absence of antibiotics (Figure 4). Our representative results show the evolution of populations at 37 &#176;C and a dilution rate of 10-4. Following the abundance of plasmid-carrying cells, we observed a decrease in the frequency plasmid-carrying host cells over time (Figure 4). Our approach enabled us to discover that the plasmid loss was a result of the condensed plasmid genome architecture, which led to plasmid instability caused by conflicts introduced by transcription of the resistance gene and replication of the plasmid itself. Visualizing the plasmid molecules enabled us to discover that these conflicts led to an unstable plasmid conformation (i.e., plasmid multimer formation, Figure 5). Nonetheless, we observed plasmid stability evolution without the exposure to antibiotics (Figure 4). The evolved stability was conferred by a plasmid intrinsic duplication that abolished the transcription-replication conflicts and led to the formation of stably inherited plasmids. Our results thus demonstrate the importance of recombination and genome amplification in adaptive evolution of genetic elements.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60749/60749fig1.jpg" /> 
Figure 1: Plasmid design of pCON. Schematic representation of the cloning strategy used to build the plasmid pCON. The plasmid backbone (pBBR1) and antibiotic resistance gene (nptII) are PCR amplified and fused by isothermal fusion20. This yields the plasmid pCON and the strain MG1655 pCON. <a href="https://www.jove.com/files/ftp_upload/60749/60749fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60749/60749fig2.jpg" /> 
Figure 2: Design of the long-term evolution experiment. Schematic representation of the serial transfer experiment. The plasmid-carrying (pCON) populations are plated on selective media. Ancestral colonies are randomly chosen from the plate and introduced to a serial transfer system. The transfers are conducted with three different dilution approaches to simulate population bottlenecks of different sizes. The dilutions are serially repeated. The experiment is conducted in two temperature regimes. The plasmid-host frequency is measured along the experiment via replica plating. <a href="https://www.jove.com/files/ftp_upload/60749/60749fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60749/60749fig3.jpg" /> 
Figure 3: Replica plating. Schematic representation of the steps used in replica plating of bacterial populations. <a href="https://www.jove.com/files/ftp_upload/60749/60749fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60749/60749fig4.jpg" /> 
Figure 4: Representative pCON frequency over time. pCON persistence is shown as the proportion of hosts (representative replicate populations) during the evolution experiment. For 98 transfers, pCON plasmid populations evolved under nonselective conditions with a dilution factor of 10-4. All replicates carrying the plasmid pCON decreased in the population. Afterwards, the populations were exposed to antibiotics for overnight incubation and were cultivated again under nonselective conditions to test for plasmid stability evolution. This figure has been modified from Wein et al.8 <a href="https://www.jove.com/files/ftp_upload/60749/60749fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60749/60749fig5.jpg" /> 
Figure 5: Representative analysis of the plasmid conformation. Visualization of the model plasmid pCON. Visualized is the untreated plasmid DNA directly after extracting, linearized plasmid DNA, and treated with DNase that only cuts chromosomal DNA as well as enzymatically nicked DNA (i.e., open circle plasmid DNA). Linearizing the plasmid shows that all plasmids are of the same size. Removing chromosomal DNA and nicking pCON reveals the presence of dimers and other multimers. This figure has been modified from Wein et al.8. <a href="https://www.jove.com/files/ftp_upload/60749/60749fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Quantification of Plasmid-Mediated Antibiotic Resistance in an Experimental Evolution Approach at JoVE.com

Quantification of Plasmid-Mediated Antibiotic Resistance in an Experimental Evolution Approach

Our experimental approach provides a strategy to follow plasmid abundance and antibiotic resistance over time in bacterial populations.

Plasmids play a major role in microbial ecology and evolution as vehicles of lateral gene transfer and reservoirs of accessory gene functions in microbial ...

quantification-plasmid-mediated-antibiotic-resistance-an-experimental

Research

JoVE Journal

Genetics

13.8K Views.  Kiel University. Our experimental approach provides a strategy to follow plasmid abundance and antibiotic resistance over time in bacterial populations.

Video: Quantification of Plasmid-Mediated Antibiotic Resistance in an Experimental Evolution Approach

Design and Use of a Low Cost, Automated Morbidostat for Adaptive Evolution of Bacteria Under Antibiotic Drug Selection

We describe a low cost, configurable morbidostat for characterizing the evolutionary pathway of antibiotic resistance. The morbidostat is a bacterial culture device that continuously monitors bacterial growth and dynamically adjusts the drug concentration to constantly challenge the bacteria as they evolve to acquire drug resistance. The device features a working volume of ~10 ml and is fully automated and equipped with optical density measurement and micro-pumps for medium and drug delivery. To validate the platform, we measured the stepwise acquisition of trimethoprim resistance in Escherichia coli MG 1655, and integrated the device with a multiplexed microfluidic platform to investigate cell morphology and antibiotic susceptibility. The approach can be up-scaled to laboratory studies of antibiotic drug resistance, and is extendible to adaptive evolution for strain improvements in metabolic engineering and other bacterial culture experiments.

We describe a low cost, configurable morbidostat that enables the characterization of antibiotic drug resistance by dynamically adjusting the drug concentration. The device can be integrated with a multiplexed microfluidic platform. The approach can be scaled up for laboratory antibiotic drug resistance studies.

We describe a low cost, configurable morbidostat for characterizing the evolutionary pathway of antibiotic resistance. The morbidostat is a bacterial ...

Testing the Role of Multicopy Plasmids in the Evolution of Antibiotic Resistance

Multicopy plasmids are extremely abundant in prokaryotes but their role in bacterial evolution remains poorly understood. We recently showed that the increase in gene copy number per cell provided by multicopy plasmids could accelerate the evolution of plasmid-encoded genes. In this work, we present an experimental system to test the ability of multicopy plasmids to promote gene evolution. Using simple molecular biology methods, we constructed a model system where an antibiotic resistance gene can be inserted into Escherichia coli MG1655, either in the chromosome or on a multicopy plasmid. We use an experimental evolution approach to propagate the different strains under increasing concentrations of antibiotics and we measure survival of bacterial populations over time. The choice of the antibiotic molecule and the resistance gene is so that the gene can only confer resistance through the acquisition of mutations. This &#34;evolutionary rescue&#34; approach provides a simple method to test the potential of multicopy plasmids to promote the acquisition of antibiotic resistance. In the next step of the experimental system, the molecular bases of antibiotic resistance are characterized. To identify mutations responsible for the acquisition of antibiotic resistance we use deep DNA sequencing of samples obtained from whole populations and clones. Finally, to confirm the role of the mutations in the gene under study, we reconstruct them in the parental background and test the resistance phenotype of the resulting strains.

Here we present an experimental method to test the role of multicopy plasmids in the evolution of antibiotic resistance.

Multicopy plasmids are extremely abundant in prokaryotes but their role in bacterial evolution remains poorly understood. We recently showed that the ...

Adherence of Bacteria to Plant Surfaces Measured in the Laboratory

This manuscript describes a method to measure bacterial binding to axenic plant surfaces in the light microscope and through the use of viable cell counts. Plant materials used include roots, sprouts, leaves, and cut fruits. The methods described are inexpensive, easy, and suitable for small sample sizes. Binding is measured in the laboratory and a variety of incubation media and conditions can be used. The effect of inhibitors can be determined. Situations that promote and inhibit binding can also be assessed. In some cases it is possible to distinguish whether various conditions alter binding primarily due to their effects on the plant or on the bacteria.

An easy method for measuring and characterizing bacterial adhesion to plants, particularly roots and sprouts, is described in this article.

This manuscript describes a method to measure bacterial binding to axenic plant surfaces in the light microscope and through the use of viable cell ...

Environment

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

A first approach to study the function of an unknown gene in bacteria is to create a knock-out of this gene. Here, we describe a robust and fast protocol for transferring gene deletion mutations from one Escherichia coli strain to another by using generalized transduction with the bacteriophage P1. This method requires that the mutation be selectable (e.g., based on gene disruptions using antibiotic cassette insertions). Such antibiotic cassettes can be mobilized from a donor strain and introduced into a recipient strain of interest to quickly and easily generate a gene deletion mutant. The antibiotic cassette can be designed to include flippase recognition sites that allow the excision of the cassette by a site-specific recombinase to produce a clean knock-out with only a ~100-base-pair-long scar sequence in the genome. We demonstrate the protocol by knocking out the tamA gene encoding an assembly factor involved in autotransporter biogenesis and test the effect of this knock-out on the biogenesis and function of two trimeric autotransporter adhesins. Though gene deletion by P1 transduction has its limitations, the ease and speed of its implementation make it an attractive alternative to other methods of gene deletion.

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

Here we present a protocol for the use of pre-existing antibiotic resistance-cassette deletion constructs as a basis for making deletion mutants in other E. coli strains. Such deletion mutations can be mobilized and inserted into the corresponding locus of a recipient strain using P1 bacteriophage transduction.

A first approach to study the function of an unknown gene in bacteria is to create a knock-out of this gene. Here, we describe a robust and fast protocol ...

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

Biology

Single&#45;Copy Gene Expression for Characterizing Multidrug Efflux Systems

Characterizing Multidrug Efflux Systems in Acinetobacter baumannii Using an Efflux&#45;Deficient Bacterial Strain and a Single&#45;Copy Gene Expression System

Acinetobacter baumannii is recognized as a challenging Gram-negative pathogen due to its widespread resistance to antibiotics. It is crucial to comprehend the mechanisms behind this resistance to design new and effective therapeutic options. Unfortunately, our ability to investigate these mechanisms in A. baumannii is hindered by the paucity of suitable genetic manipulation tools. Here, we describe methods for utilizing a chromosomal mini-Tn7-based system to achieve single-copy gene expression in an A. baumannii strain that lacks functional RND-type efflux mechanisms. Single-copy insertion and inducible efflux pump expression are quite advantageous, as the presence of RND efflux operons on high-copy number plasmids is often poorly tolerated by bacterial cells. Moreover, incorporating recombinant mini-Tn7 expression vectors into the chromosome of a surrogate A. baumannii host with increased efflux sensitivity helps circumvent interference from other efflux pumps. This system is valuable not only for investigating uncharacterized bacterial efflux pumps but also for assessing the effectiveness of potential inhibitors targeting these pumps.

Author Spotlight: Advancing Antibiotic Resistance Research Using an Efflux&#45;Deficient Bacterial Strain and a Single&#45;Copy Gene Expression System

We describe a facile procedure for the single-copy chromosomal complementation of an efflux pump gene using a mini-Tn7-based expression system into an engineered efflux-deficient strain of Acinetobacter baumannii. This precise genetic tool allows for controlled gene expression, which is key for the characterization of efflux pumps in multidrug resistant pathogens.

Acinetobacter baumannii is recognized as a challenging Gram-negative pathogen due to its widespread resistance to antibiotics. It is crucial to comprehend ...

Immunology and Infection

Plasmid Stability Analysis with Open-Source Droplet Microfluidics

Plasmids play a vital role in synthetic biology by enabling the introduction and expression of foreign genes in various organisms, thereby facilitating the construction of biological circuits and pathways within and between cell populations. For many applications, maintaining functional plasmids without antibiotic selection is critical. This study introduces an open-hardware-based microfluidic workflow for analyzing plasmid retention by culturing single cells in gel microdroplets and quantifying microcolonies using fluorescence microscopy. This approach allows for the parallel analysis of numerous droplets and microcolonies, providing greater statistical power compared to traditional plate counting and enabling the integration of the assay into other droplet microfluidic workflows. By using plasmids expressing fluorescent proteins alongside a non-specific fluorescent DNA stain, single colonies can be identified and differentiated based on plasmid loss or fluorescent marker expression. Notably, this advanced workflow, implemented with open-source hardware, offers precise flow control and temperature management of both the sample and the microfluidic chip. These features enhance the workflow&#39;s ease of use, robustness, and accessibility. While the study focuses on Escherichia coli as the experimental model, the method&#39;s true potential lies in its versatility. It can be adapted for various studies requiring fluorescence signal quantification from plasmids or stains, as well as for other applications. The adoption of open-source hardware broadens the potential for conducting high-throughput bioanalyses using accessible technology in diverse research settings.

An accessible, open-source microfluidic workflow is presented for the parallelized analysis of plasmid retention in bacteria. By employing fluorescence microscopy to quantify plasmid presence in single-cell microcolonies within gel microdroplets, this method provides a precise, accessible, and scalable alternative to traditional plate counting.

Plasmids play a vital role in synthetic biology by enabling the introduction and expression of foreign genes in various organisms, thereby facilitating ...

Bioengineering

A Protocol for Functional Assessment of Whole-Protein Saturation Mutagenesis Libraries Utilizing High-Throughput Sequencing

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel sequencing technology have opened up the possibility of assessing the functional or fitness effects of large numbers of mutations simultaneously. Here, we present a protocol for experimentally determining the effects of all possible single amino acid mutations in a protein of interest utilizing high-throughput sequencing technology, using the 263 amino acid antibiotic resistance enzyme TEM-1 &#946;-lactamase as an example. In this approach, a whole-protein saturation mutagenesis library is constructed by site-directed mutagenic PCR, randomizing each position individually to all possible amino acids. The library is then transformed into bacteria, and selected for the ability to confer resistance to &#946;-lactam antibiotics. The fitness effect of each mutation is then determined by deep sequencing of the library before and after selection. Importantly, this protocol introduces methods which maximize sequencing read depth and permit the simultaneous selection of the entire mutation library, by mixing adjacent positions into groups of length accommodated by high-throughput sequencing read length and utilizing orthogonal primers to barcode each group. Representative results using this protocol are provided by assessing the fitness effects of all single amino acid mutations in TEM-1 at a clinically relevant dosage of ampicillin. The method should be easily extendable to other proteins for which a high-throughput selection assay is in place.

We present a protocol for the functional assessment of comprehensive single-site saturation mutagenesis libraries of proteins utilizing high-throughput sequencing. Importantly, this approach uses orthogonal primer pairs to multiplex library construction and sequencing. Representative results using TEM-1 &#946;-lactamase selected at a clinically relevant dosage of ampicillin are provided.

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel ...

Population and Single-Cell Analysis of Antibiotic Persistence in Escherichia coli

Antibiotic persistence refers to the capacity of small bacterial subpopulations to transiently tolerate high doses of bactericidal antibiotics. Upon bactericidal antibiotic treatment, the bulk of the bacterial population is rapidly killed. This first rapid phase of killing is followed by a substantial decrease in the rate of killing as the persister cells remain viable. Classically, persistence is determined at the population level by time/kill assays performed with high doses of antibiotics and for defined exposure times. While this method provides information about the level of persister cells and the killing kinetics, it fails to reflect the intrinsic cell-to-cell heterogeneity underlying the persistence phenomenon. The protocol described here combines classical time/kill assays with single-cell analysis using real-time fluorescence microscopy. By using appropriate fluorescent reporters, the microscopy imaging of live cells can provide information regarding the effects of the antibiotic on cellular processes, such as chromosome replication and segregation, cell elongation, and cell division. Combining population and single-cell analysis allows for the molecular and cellular characterization of the persistence phenotype.

Population and Single-Cell Analysis of Antibiotic Persistence in Escherichia coli

Antibiotic persistence describes the ability of small subpopulations within a sensitive isogenic population to transiently tolerate high doses of bactericidal antibiotics. The present protocol combines approaches to characterize the antibiotic persistence phenotype at the molecular and cellular levels after exposing Escherichia coli to lethal doses of ofloxacin.

Antibiotic persistence refers to the capacity of small bacterial subpopulations to transiently tolerate high doses of bactericidal antibiotics. Upon ...

Phage Transduction: A Method to Transfer Ampicillin Resistance from Donor to Recipient E. coli

Source: Alexander S. Gold1, Tonya M. Colpitts1 
1 Department of Microbiology, Boston University School of Medicine, National Emerging Infections Diseases Laboratories, Boston, MA
Transduction is a form of genetic exchange between bacteria that utilizes bacteriophages, or phages, a class of virus that infects exclusively prokaryotic organisms. This form of DNA transfer, from one bacterium to another by way of a phage, was discovered in 1951 by Norton Zinder and Joshua Ledererg, who termed the process &#34;transduction&#34; (1). Bacteriophages were first discovered in 1915 by British bacteriologist Frederick Twort, then independently discovered again in 1917 by French-Canadian microbiologist Felix d&#39;Herelle (2). Since then, the structure and function of these phages have been widely characterized (3), dividing these phages into two classes. The first of these classes are the lytic phages which upon infection multiply within the host bacterium, disrupting the bacterial metabolism, lysing the cell, and releasing progeny phage (4). As a result of this anti-bacterial activity and the increasing prevalence of antibiotic-resistant bacteria, these lytic phages have recently proved useful as a substitute treatment for antibiotics. The second of these classes are the lysogenic phages which can either multiply within the host via the lytic cycle or enter a quiescent state in which their genome is integrated into that of the host (Figure 1), a process known as lysogeny, with the ability for phage production to be induced in multiple later generations (4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10517/10517fig1.jpg" /> 
Figure 1: Infection of host cell by bacteriophage. Adsorption by the phage to the bacterial cell wall via interactions between the tail fibers and receptor (purple). Once on the cell surface, the phage is irreversibly attached to the bacterial cell using the base plate (black) which is moved to the cell wall by the contractile sheath (yellow). Phage genome (red) then enters the cell and integrates into the host cell genome.
While bacterial transduction is a naturally occurring process, using modern technology it has been manipulated for the transfer of genes into bacteria in the laboratory setting. By inserting genes of interest into the genome of a lysogenic phage, such as phage, one is able to transfer these genes into the genomes of bacteria and consequently express them within these cells. While other methods of gene transfer, such as transformation, use a plasmid for gene transfer and expression, the insertion of the phage genome into that of the recipient bacterium not only has the potential to confer new traits to this bacterium, but also allows for naturally occurring mutations and other factors of the cellular environment to alter the function of the transferred gene.
Compared to other methods of horizontal gene transfer, such as conjugation, transduction is fairly flexible in the criteria required for donor and recipient cells. Any genetic element that can fit inside the genome of the phage being used can be transferred from any strain of donor bacteria to any strain of recipient bacteria as long as both are permissive to the phage, requiring the expression of necessary phage receptors on the cell surfaces. Once this gene is moved out of the donor genome and packaged into the phage, it can be transferred to the recipient. Following transduction, it is necessary to select for recipient bacteria that contain the gene of interest have to be selected for. This could be done by use of a genetic marker, such as a FLAG-tag or polyhistidine-tag, to mark the gene of interest, or the intrinsic function of the gene, in the case of genes that encode for antibiotic resistance. Additionally, PCR could be used to further confirm successful transduction. By using primers for a region within the gene of interest and comparing signal to a positive control, bacteria that has the gene of interest, and a negative control, bacteria that underwent the same steps as the transduction reaction with no phage. While bacterial transduction is a useful tool in molecular biology, it has and continues to play an important role in the evolution of bacteria, particularly with regard to the recent rise of antibiotic resistance.
In this experiment, bacterial transduction was used to transfer the gene encoding for resistance to the antibiotic ampicillin from the W3110 strain of E.coli to the J53 strain via the P1 bacteriophage (5). This experiment consisted of two main steps. First, the preparation of P1 phage containing the ampicillin resistance gene from the donor strain. Second, the transfer of this gene to the recipient strain by transduction with the P1 phage (Figure 1). Once carried out, the successful transfer of the ampicillin resistance gene could be determined by qPCR (Figure 2). If transduction was successful, the J53 strain of E. coli would be resistant to ampicillin, and the gene conferring this resistance detectable by qPCR. If unsuccessful, there would be no detection of the ampicillin resistance gene and ampicillin would still function as an effective antibiotic against the J53 strain.

<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/10517/10517fig2.jpg" /> 
Figure 2: The confirmation of successful transduction by qPCR. By comparing the Cq values detected for the gene of interest from the transduction reaction and the negative control reaction, and normalizing these values against a housekeeping gene, one was able to confirm that bacterial transduction was successful.

Phage Transduction: Transferring Ampicillin Resistance in E. coli

Source: Alexander S. Gold1, Tonya M. Colpitts1
1 Department of Microbiology, Boston University School of Medicine, National Emerging Infections Diseases ...

Quantification of Plasmid-Mediated Antibiotic Resistance in an Experimental Evolution Approach

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Design and Use of a Low Cost, Automated Morbidostat for Adaptive Evolution of Bacteria Under Antibiotic Drug Selection

Testing the Role of Multicopy Plasmids in the Evolution of Antibiotic Resistance

Adherence of Bacteria to Plant Surfaces Measured in the Laboratory

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Characterizing Multidrug Efflux Systems in Acinetobacter baumannii Using an Efflux-Deficient Bacterial Strain and a Single-Copy Gene Expression System

Plasmid Stability Analysis with Open-Source Droplet Microfluidics

A Protocol for Functional Assessment of Whole-Protein Saturation Mutagenesis Libraries Utilizing High-Throughput Sequencing

Population and Single-Cell Analysis of Antibiotic Persistence in Escherichia coli

Phage Transduction: A Method to Transfer Ampicillin Resistance from Donor to Recipient E. coli