High-throughput Parallel Sequencing to Measure Fitness of Leptospira interrogans Transposon Insertion Mutants During Golden Syrian Hamster Infection

The overall goal of this high-throughput transposon sequencing technique is to identify and quantify transposon mutants with in vivo and in vitro fitness defects, and thereby, identify Leptospira genes involved in virulence. This method can help answer key questions in the Leptospira field, such as identifying genes involved in colonization, dissemination, and survival of the bacteria in the host. The main advantage of this technique is that a large number of mutants can be screened with a limited number of animals.

Though this method was designed to examine the fitness of Leptospira interrogans mutants in the hamster model with acute leptospirosis, it can also be applied through a chronic infection models, in vitro studies, and other strains of Leptospira. Demonstrating the procedure will be Kritel Lourdault and Karen Evangelista, both post-docs from my laboratory. First, assemble a filtration unit.

Place the base on a 125 milliliter side arm Erlenmeyer flask. Position an acetate cellulose filter at the base and clamp a funnel onto the base. Connect the filtration unit to a vacuum system.

Next, add five milliliters of Leptospira culture and 0.5 milliliters of E.coli culture into the funnel. Turn on the vacuum to filter the solution. Then place the filter with bacteria-side up onto an EMJH plate, supplemented with DAP.

Incubate the plate at 30 degrees Celsius over night. The next day, transfer the filter into a 15 milliliter tube containing one milliliter of EMJH, and vortex the tube for 10 seconds to release the bacteria in the suspension. Then spread the suspension onto five EMJH plates, supplemented with kanamycin using glass beads or a spreader.

Wrap the plates in parafilm and incubate them upside down at 30 degrees Celsius until colonies are visible. When colonies are visible, collect and transfer them individually into three milliliters of EMJH containing kanamycin. Then incubate the tubes at 30 degrees Celsius with 150 rpm agitation for seven to 10 days, or to a density of 100 million cells per milliliter.

To identify the transposon insertion site, perform nested PCRs. First, prepare the mix for the first PCR. Then, add the samples from culture to the PCR mixes.

After running the first reaction, prepare the mix for the second PCR. Add two microliters of PCR product from the first PCR to the complete the reaction mixes. After running the PCR, examine the products on an agarose gel.

Proceed by sequencing the products to determine the transposon insertion site. To begin the animal experiment, first grow individual mutants in EMJH, supplemented with kanamycin. Then determine the culture densities using a dark field microscope.

Dilute each cultured mutant strain in EMJH to the same density. Then, pool an equal volume of all the mutants to make the input pool. Now challenge the animals as described in the text protocol.

The number of animals required depends on the size of the input pool. Be sure to consult a biostatistician to determine the number of animals to inoculate. Allow the infection to develop for four days, then harvest blood and tissues as described in the text protocol.

To extract the DNA from the blood sample, use a commercial extraction kit. For tissue, dice 50 to 80 milligrams of tissue into one millimeter cubes. Then, transfer the diced tissues to a screw cap tube and weigh them on a precision balance.

Next, add 500 microliters of sterile PBS into the tube. Homogenize it using a disrupter for one minute. Then use a volume corresponding to 25 milligrams of tissue to extract the DNA.

Use a commercial kit according to the manufacturer's instructions. Dilute the DNA in 100 microliters of elution buffer. Next, transfer 50 microliters of extracted DNA into a 1.5 milliliter microcentrifuge tube.

Then shear the DNA. Place the tube into a sonicator cup horn filled with water at four degrees Celsius. Run the sonicator for three minutes while wearing ear protection.

Next, load 2.5 microliters of the sheared DNA into a 2%agarose gel to confirm that most of the DNA fragments are under 600 base pairs. Next, prepare the reaction mix to add cytosine tails to the DNA fragments. Incubate the reaction for an hour at 37 degrees Celsius.

Then, inactivate the enzyme by increasing the temperature to 75 degrees Celsius for 20 minutes. When the reaction is done, clean it up using a PCR purification kit and elute the DNA with 12 microliters of elution buffer. Run two PCRs to create the genomic libraries.

For the first PCR, prepare mixes with TnkN3 and olj376 primers. Primer TnkN3 is specific to the transposon and primer olj376 is specific to the C-tail. For each sample, combine 22 microliters of mix with three microliters of C-tailed purified DNA.

Then, run the PCR as described in the text protocol. For the second PCR, use one microliter of the first PCR product as a template. For primers, use pMargent2 to target the transposon and use the IP primers.

The IP primers each have a six base pair barcode and specific sequences recognizable by the next generation sequencing platform. Next run out three microliters of the second PCR products on a 2%agarose gel. The library should show a smear with the majority of the signal between 200 to 600 base pairs, and no amplification should be observable in the control reaction.

Next, clean the genomic libraries with a PCR purification kit and elute the DNA with 30 microliters of elution buffer. Then measure the DNA concentration with a fluorometer. Now, calculate the volume of each library needed for 15 to 20 nanograms of DNA.

Mix the libraries and determine the DNA molar concentration in the pool, using a publicly available calculator. Now send the pooled genomic libraries out for high-throughput sequencing. Later, process the FASTQ files obtained from sequencing using open source software and analyze the results.

Eight hamsters were challenged with one milliliter of pooled transposon mutants and the infection was allowed to develop for four days. After extracting the DNA collected from these hamsters'livers, it was sheared and characterized on an agarose gel. Most of the fragments were between 200 and 600 base pairs.

After adding C-tail to the sheared DNA, PCRs were used to prepare genomic libraries. Electrophoresis was performed to confirm the size of the PCR products. The size of most of the PCR products was between 200 and 600 base pairs.

After processing the sequencing reads, the output/input ratios were determined using the transposon sequencing approach. As expected, mutants known to be virulent, such as flaA1 and IlgB do not show any change in fitness. In contrast, mutants known to be attenuated, such as loa22 show a decrease in fitness.

Mutants with a reduced in vivo fitness level were identified, such as lic12327. Follow up experimentation showed the virulence of these mutants to be attenuated in hamsters. After watching this video, you should have a good understanding of how to generate genomic DNA libraries in order to quantify and identify mutants with altered in vivo fitness.

After mutants with in vivo fitness defects are identified, they can be tested individually in hamsters to determine if their virulence is attenuated. Don't forget that Leptospira interrogans can be lethal and must be handled using BSL2 containment. Personal protective equipment including gloves and a lab coat must be worn while handling the bacteria.