Identification of Virulence Markers of Mycobacterium abscessus for Intracellular Replication in Phagocytes

1. Library Screening

1. Construction of the Tn mutant library
   1. Obtain a transposon library.
      NOTE: For this experiment, a transposon mutant library was obtained from E.J. Rubin, Harvard School of Public Health, Boston, USA. The library was constructed from a smooth clinical strain (43S) of the M. abscessus complex (M. abscessus subsp. massiliense) with a phagemid introduced into M. abscessus allowing the random insertion of a single Tn into a TA dinucleotide (91,240 TA sequences in the M. abscessus genome). For more details see reference of Rubin et al.⁹ and Laencina et al.¹⁰.
2. Titration of the library and conservation of the M. abscessusTn mutants in plates
   NOTE: This takes one week.
   1. Thaw the library on the ice. Determine the bacterial concentration by performing serial dilutions in 1 mL of water (10^-1 to 10^-15). Plate a drop of each dilution on a Petri dish containing 7H11 agar medium with 250 mg/mL kanamycin. Incubate the plates at 37 °C for 3–4 days.
      NOTE: Here, a titration of 10¹¹ bacteria/mL were recovered.
   2. After determining the concentration of the library, dilute the library to a final concentration of 3,000 bacteria/mL in 10 mL of 25% glycerol-water. Aliquot the library in 10 cryotubes with 1 mL of the library in each tube. Store the aliquots at -80 °C.
   3. When ready to use, thaw one aliquot of the diluted library on ice and add 100 µL of the library to 10 square Mueller-Hinton (MH) agar plates (size 12 cm) to recover 200 to 300 individual colonies after 3–4 days of incubation at 37 °C.
   4. With sterile toothpicks, transfer the individual colonies (about 6,000 colonies in 60 x 96-well plates) into 96-well plates filled with 200 µL of 7H9 medium supplemented with 0.2% glycerol, 1% glucose, 250 mg/mL of kanamycin. Incubate at 37 °C for 5 days without shaking.
   5. Transfer 100 µL of the culture (step 1.2.4) to a 96-well plate containing 100 µL of 7H9 medium with 40% glycerol. Store the ordered library at -80 °C.
   6. Repeat steps 1.2.3. to 1.2.5. with the rest of the library until 10,000 individual clones are recovered (around 100 x 96-well plate and 30 MH plates).
3. Preparation of amoebae
   1. Culture amoeba Acanthamoeba castellanii (AC) in 75 cm² tissue culture flasks at room temperature (RT) in 30 mL of PYG medium (Table 1) for amplification of the cell line¹¹.
      NOTE: Mature trophozoites from large adhesive cells with numerous digestive vacuoles are clearly visible on a microscope. The cell doubling time is estimated to be 25 h. Cells were amplified every 3 days by spreading one third of the initial culture in 75 cm² tissue culture flasks.
   2. Remove the PYG medium with a 25 mL pipette. Wash the cells with 10 mL of AC buffer (Table 2), which does not contain any carbon or nitrogen sources¹². Repeat the wash twice more.
      NOTE: AC buffer that contains no carbon or nitrogen sources avoids the extracellular growth of mycobacteria. This minimum medium leads to the encystment of amoebae. To limit this, heat-inactivated E. coli were added as a substrate (step 1.4) to amoebae and very short culture times were promoted (48 h for mutant screening and 4 h or 16 h culture for RNAseq experiments). Alternatively, use this medium but enriched with PYG medium (usually 10%).
   3. Detach the amoeba from the bottom of the flask with a single vigorous shake of the tissue culture flask.
   4. Count cells with a hemocytometer to adjust the cell concentration to 5 x 10⁵ amoebae/mL diluted in AC buffer.
4. Preparation of heat-inactivated Escherichia coli bacteria to feed amoeba after infection
   1. Grow the E. coli clinical isolate strain from a glycerol stock in 100 mL of Luria Broth (LB) overnight at 37 °C.
   2. Harvest the bacteria by centrifugation at 1,835 x g for 10 min in 50 mL tubes. Discard the supernatant. Resuspend the pellet with 10 mL of 10% glycerol-water. Repeat the washing twice more.
   3. Resuspend the pellet with 5 mL of 10% glycerol-water. Aliquot 0.5 mL into 1.5 mL tubes. Determine the amount of bacteria/mL by performing serial dilutions and adding dilutions on LB agar plates. Store the aliquots at -80 °C.
   4. The day before the amoeba infection, thaw one aliquot on ice and proceed to heat-killing by incubating the tube at 70 °C for 60 min.
      NOTE: Check the inactivation by plating 100 µL of inactivated bacteria on LB agar plates overnight at 37 °C. No colonies should be visible after one night of culture.
   5. Dilute the heat-killed bacteria to a concentration of 10⁷ bacteria in 50 µL of AC buffer and store the diluted bacteria at 4 °C for no more than one week.
5. Co-culture of amoebae-M. abscessus Tn mutant: first screen
   NOTE: This takes 3–4 months for the screening of 6,000 mutants.
   1. Spread 5 x 10⁴ amoebae/well to a 96-well plate in 100 µL of AC buffer. Incubate for 1 h at 32 °C without shaking. Allow the floating amoeba trophozoïtes time to adhere to the wells.
      1. While incubating, thaw bacteria on ice for 1 h.
   2. Add 5 µL of the thawed bacteria cultures with an electronic multichannel pipette. Infect for 1.5 h. Screen around 6,000 individualized clones from the 96-well plate Tn mutants stock.
      NOTE: The MOI is unknown at this step of the protocol; however, all the 96-well plates containing Tn mutants were cultivated in exactly the same way. This first screen aims to quickly identify intracellular deficient mutants, without considering the variations in inoculum density.
   3. After the 1.5 h of infection, invert the 96-well plate on sterile gauzes placed in a sterilized metal tray to remove the AC medium under a fume hood. Refill with 200 µL of AC medium. Repeat this wash twice more.
   4. Add 200 µL of fresh medium supplemented with 100 µg/mL amikacin to eliminate remaining extracellular mycobacteria.
   5. Incubate for 2 h before proceeding with 3 additional washings (as described in step 1.5.3). After washing, maintain the cultures with 50 µg/mL amikacin in 200 µL of AC buffer.
   6. Add 5 x 10⁷ heat-inactivated Escherichia coli bacteria (from step 1.4) at the end of the infection and every 24 h to prevent encystment of amoebae.
   7. After 48 h of co-culture, lyse the cells by adding 10 µL of 10% SDS (sodium dodecyl sulfate) to each well and incubate for 30 min at 32 °C. Proceed with the serial dilution and add on Columbia agar plates containing 5% sheep blood (COS plates) to evaluate the number of intracellular mycobacteria. Seal the plate with the parafilm and incubate at 37 °C.
   8. When colonies appear on the agar plates after 3-4 days, photograph the plate and identify the mutants impaired for intracellular growth by observing the deposits with the lowest number of bacterial colonies.
      NOTE: Do not mind about the shape, height or density of the colony (Figure 1A). 136/6000 mutants were identified.
6. Co-culture of amoebae-M. abscessusTn mutant: second screen of the mutants identified during the first screen
   NOTE: This takes 2 weeks. A second screen must be performed with the 136 attenuated mutants obtained after the first screen to quantify precisely the number of bacteria inoculated and the number of intracellular survival bacteria 2 days post-infection.
   1. Grown 1 colony of each impacted mutant in 50 mL tubes filled with 20 mL of 7H9 medium supplemented with 0.2% glycerol, 1% glucose and 250 mg/mL of kanamycin. Allow the bacteria to reach the exponential phase (0.6<OD<0.8).
   2. Wash the culture by centrifugation (1,835 x g for 10 min) in AC medium. Discard the supernatant. Repeat this wash twice more with 20 mL of 7H9 medium supplemented with 0.2% glycerol, 1% glucose and 250 mg/mL of kanamycin.
   3. Resuspend the bacteria in 5 mL of AC buffer.
   4. Determine the colony forming unit (CFU) concentration of the mutants. In 24-well plates, add 1 mL of water to the two first rows. Add 100 µL of the bacterial suspension to the first well. With a micropipette, mix three times. Then, aspirate 100 µL and deposit to the second well.
   5. With a new tip, repeat step 1.6.4 from the second well to the third one, etc. until the twelfth well.
   6. Aspirate 30 µL of the 10^-12 dilution and add it to a COS plate. Repeat for the other dilutions.
   7. Wrap the COS plate with parafilm and incubate for 3–4 days at 37 °C. Determine the concentration by counting the colonies.
   8. Perform co-culture assays as described above in triplicate in a 24-well plate with 5 x 10⁴ amoeba per well in 1 mL of AC co-culture medium. Inoculate with a multiplicity of infection (MOI) of 10.
   9. After 48 h of co-culture, proceed with the cell lysis as described in step 1.5.7. Perform serial dilutions in water (10^-1 to 10^-6) and add 30 µL to the COS plates. Incubate the plates for 4 days at 37 °C before proceeding with CFU counting.
      NOTE: After this second screen, 60 of 136 mutants were conserved.
   10. Store the mutants impacted for their survival inside the cells. Add a colony to a cryotube containing LB medium with glycerol (20% final) or in a cryotube containing commercial solution and microbeads to favor a long-term conservation and facilitate future inoculation and then store at -80 °C.
   11. Assess the in vitro mutant growth by measuring the optical density (OD) at 600 nm every 2 days.
       1. Grow 1 colony of each impacted mutant in 50 mL tubes filled with 20 mL of 7H9 medium supplemented with 0.2% glycerol, 1% glucose and 250 mg/mL of kanamycin. Allow the bacteria to reach the exponential phase (0.6<OD<0.8) before adjusting the OD to 0.01 to begin the kinetics.
       2. Exclude mutants with an in vitro growth defect when compared with the WT strain from the set of intracellular deficient mutants.
7. Co-culture of macrophages-M. abscessus Tn mutant defective for an intra-amoeba life
   NOTE: This takes 1 month.
   1. Spread 5 x 10⁴ J774.2 murine macrophages per well of a 24-well plate in 1 mL of rich medium, DMEM supplemented with 10% of Fetal Calf Serum (FCS). Incubate the plates for 24 h at 37 °C and 5% CO₂ to allow macrophage adhesion. Provide 3 replicates.
   2. Perform the infection. Inoculate Tn mutants, with a MOI of 10, diluted in DMEM with 10% FCS in a volume of 100 µL.
   3. After 3 h of infection, perform three washings with 1 mL of DMEM (a vacuum pump can be used). Then treat with 250 µg/mL amikacin (in DMEM with 10% FCS) for 1 h to eliminate remaining extracellular mycobacteria.
      1. After 3 additional washes with 1 mL of DMEM, maintain the cultures in 250 µg/mL amikacin (in DMEM with 10% FCS) to a final volume of 1 mL to prevent extracellular mycobacterial multiplication.
   4. 72 h after co-culture, remove the medium and lyse the macrophages by adding 1 mL of cold water. Incubate for 30 min at 4 °C.
   5. Perform CFU assays on COS plates to evaluate the number of intracellular bacteria.
      1. In a 24-well plate, add 1 mL of water to the two first rows. Add 100 µL of the co-cell lysate to the first well. With a micropipette, mix three times. Aspirate 100 µL and deposit them in the second well.
      2. With a new tip, repeat the dilution from the second well to the third one, etc. until the twelfth well. Then aspirate 30 µL of the 10^-12 dilution and add it to a COS plate.
      3. Repeat for the other dilutions. Wrap the COS plate with parafilm and wait 3-4 days to observe and count the colonies.
8. Identification and sequencing of the site of insertion of the Tn de M. abscessus mutants
   NOTE: This takes 1.5 months for 60 mutants.
   1. Genomic extraction of the identified M. abscessus mutants
      1. Grow the mutants in a 50 mL tube with 20 mL of 7H9 medium supplemented with 0.2% glycerol, 1% glucose and kanamycin 250 mg/mL to exponential phase (OD₆₀₀=0.8).
      2. Harvest 10 mL of the cultures (2,396 x g, 5 min). Discard the supernatant. Suspend the bacteria in 500 µL of solution I (25% sucrose, 50 mM Tris pH 8, 10 mg/mL lysozyme, 40 mM thiourea). Transfer the solution into 2 mL tubes with screw cap and incubate for 2 h at 37 °C.
      3. Add 500 µL of solution II (25% sucrose, 50 mM Tris pH 8, 50 mM EDTA). After pipetting up and down 5-10 times, add zirconium beads until a volume of 500 µL in the tube.
      4. Proceed with cell lysis with a homogenizer (3 x 6,000 rpm, 45 s) with a 1 min incubation on ice between each round.
      5. Add 5 µL of 20 mg/mL Proteinase K (100 µg/mL final) and incubate the samples overnight at 55 °C.
      6. Add 1 mL of cold of phenol/chloroform/isoamyl alcohol (25:24:1) under a fume hood. Mix the samples by vortexing before proceeding with centrifugation at RT and 16,500 x g for 30 min.
      7. After centrifugation, recover the aqueous phase and add an equal volume of cold phenol/chloroform/isoamyl alcohol solution under a hood. Centrifuge the samples at 16,500 x g for 30 min to recover the aqueous phase again.
      8. Add 0.7 volume of RT isopropanol to the recovered aqueous phase under a fume hood. Invert slowly the tubes to allow DNA precipitation and incubate 5 min at RT. Then centrifuge for 10 min at 16,500 x g and RT.
      9. Remove the supernatant and wash the pellet with 500 µL of 70% ethanol under a hood. Centrifuge the tubes for 10 min at 16,500 x g at RT before removing the ethanol. Incubate for 10 min to allow remaining alcohol evaporation.
      10. Finally, suspend the DNA pellet in 100 µL of DNase/RNase free water. Determine DNA yield and purity DNA with a spectrophotometer.
      11. Remove 1 µg of genomic DNA and digest with 10 U of ClaI for one night for sub-cloning the Tn insertion site. Use manufacturer's protocol.
          NOTE: The ClaI enzyme is one of the most highly represented restriction enzymes in the M. abscessus genome (2,173 restriction sites). A ClaI restriction site is also found in the Tn sequence upstream of the kanamycin resistance cassette of the Tn mutant. Thus, the ClaI-mediated subcloning enables to identify the Tn insertion site.
   2. Sub-cloning in a plasmid of genomic DNA containing the Tn
      NOTE: Before proceeding with the Tn-subcloning in the 2,686 bp pUC19 ampicillin-resistant plasmid, add a ClaI restriction site in the multiple cloning site of the plasmid. The sequence of the pUC19 plasmid can be found at this link https://www.addgene.org/50005/sequences/.
      1. Follow manufacturer's protocol to digest the pUC19 plasmid with EcoRI and HindIII, which releases a fragment of 51 bp and a fragment of 2635 bp. Purify the double digested vector with a commercial PCR purification kit to eliminate the 51 bp released fragment.
      2. Mix 1 µL (concentration 25 pmol/µL) of two complementary oligonucleotides (5'AGCT TATA AGATCT TATA ATCGAT TATA3' and 5'AAT TAA TAA TCG ATT ATA AGA TCT TAT A3') of 28 bp each containing the ClaI restriction site and at the ends the HindIII and EcoRI restriction sites. Heat at 100 °C for 10 min and then cool in order to bind them.
      3. Digest the paired primers by EcoRI and HindIII according to manufacturer's protocol.
      4. Ligate 150 ng of EcoRI/HindIII-restricted pUC19 plasmid with different concentration of paired primers (50 pmol/µL, 25 pmol/µL, 2.5 pmol/µL) for 1 h at RT with 1 U of T4 DNA ligase in a final volume of 20 µL. Check the cloning by a different enzymatic digestion.
      5. Digest the modified plasmid with ClaI for 2 h at 37 °C.
      6. Ligate 50 ng of ClaI-restricted pUC19plasmid and 50 ng of genomic DNA digested by ClaI for 1 h at RT with 1 U of T4 DNA ligase in 10 µL.
      7. Proceed to E. coli electrocompetent bacteria transformation with 5-10 µL of the ligation (2500 V, 200 Ohms, 25 µF). Select the clones on LB agar plates supplemented with 50 µg/mL kanamycin and 50 µg/mL ampicillin to select the bacteria bearing plasmids with portions of Tn sequences.
      8. Grow resistant clones overnight in LB liquid medium with the appropriate antibiotic selection (50 µg/mL kanamycin and 50 µg/mL ampicillin).
      9. Extract plasmid DNA. Check the presence of the insert by enzymatic restriction and sequence the 3' end of the Tn to identify the disrupted gene with an oligonucleotide (5' TTGACGAGTTCTTCTGA 3') corresponding to the last 17 base pairs of the Tn kanamycin resistance cassette.
      10. Align the sequence against the M. abscessus GO 06 strain by performing a nucleotide blast (BLAST-N).

2. RNA Extraction of Intracellular Mycobacteria for Rnaseq Analysis

NOTE: This takes 4 months for experiments and 4 months for RNAseq analysis.

1. Co-culture of amoebae-M. abscessus in batches
   NOTE: In the following experiment, co-culture is performed in 50 mL tubes with agitation to favor bacteria to cell contacts.
   1. Grow M. abscessus in 7H9 medium supplemented with 0.2% glycerol, 1% glucose to the mid-exponential phase at 120 rpm.
   2. Wash the bacteria twice with 10 mL of AC buffer.
   3. Estimate the bacteria concentration by measuring the OD_600nm (1 OD₆₀₀=4 x 10⁸ mycobacteria). Add 8.5 x 10⁸ mycobacteria to 8.5 x 10⁶ amoebae in 10 mL of AC medium.
   4. Incubate for 1 h at 30 °C with gentle agitation (about 80 rpm).
   5. Harvest the cells by centrifuging at 253 x g for 12 min. Remove the supernatant and suspend the cells in 10 mL of AC buffer. Repeat this wash twice more.
   6. Resuspend the cells in 10 mL of AC buffer containing 50 µg/mL amikacin to eliminate extracellular bacteria and incubate for 4 h at 30 °C with gentle agitation (80 rpm). Wash the cells twice again.
2. Extraction of total RNA from intracellular M. abscessus
   NOTE: Perform steps 2.2.1–2.2.3 under the fume hood due to the use of toxic reagents.
   1. After the final washings, resuspend the infected amoebae in 4 mL of cold solution III (Table 3) with extemporaneously 280 µL of β-mercaptoethanol to disrupt amoebae. This is the guanidinium thiocyanate (GTC) step. Maintain the suspension on ice for 10 min. Centrifuge the cell lysate for 30 min at 2,396 x g and 4 °C to pellet the released intracellular bacteria.
   2. Remove the supernatant and suspend the bacterial pellet in 1 mL of cold solution III. Harvest the bacteria at 2396 x g for 30 min at 4 °C. Resuspend the pellet in 1 mL of the extraction buffer and transfer into a 2 mL screw tube containing zirconium beads (until the 500 µL gradation of the tube). Store the tubes for at least 24 h at -80 °C.
      NOTE: Storage in the lysis reagent permits the inactivation of RNases and cell dissolution.
   3. When ready for use, thaw samples on ice and lyse them with a homogenizer by performing two rounds at 6,000 rpm for 25 s, followed by one round at 6,000 rpm for 20 s with a 1 min incubation on ice between each round.
   4. To cool the sample down, incubate them on ice for 10 min. Then add 200 µL of chloroform diluted in isoamyl alcohol. After vortexing the samples for 1 min, centrifuge for 15 min at 16,500 x g and 4 °C.
   5. Add 0.8 mL of cold isopropanol to the aqueous phase and incubate the samples for 2–3 h at 20 °C. Centrifuge the samples for 35 min at 16,500 x g and 4 °C. Discard the supernatant.
   6. Wash the RNA pellet by adding 500 µL of cold 70% ethanol (do not mix).
   7. Centrifuge the samples at 16,500 x g for 10 min to remove the ethanol. Aspirate the solution and dry in a centrifugal concentrator.
   8. Once dried, resuspend the pellets in 100 µL of DNase/RNase free water.
      NOTE: Samples must be treated twice with DNases to remove DNA contaminants according to the manufacturer's recommendations.
   9. Assess the RNA integrity on an agarose gel and confirm by measuring the RNA integrity number (RIN) and the ribosomal ratio with a chip-based capillary electrophoresis method.
      NOTE: The RIN takes the entire electrophoretic trace into account. The RIN software algorithm allows for the classification of total RNA, based on a numbering system from 1 to 10, with 1 being the most degraded profile and 10 being the most intact. To proceed with cDNA library preparation, the RINs must be over 8 and the ribosomal ratio [28S/18S] as high as possible, the ideal value being 2, depending of the organism and its specificities.
   10. Store RNA samples at -80 °C until preparation of the cDNA library for sequencing.
3. Preparation of the cDNA library
   1. To proceed with library preparation, use a commercially available cDNA synthesis kit (Table of Materials) using the manufacturer's protocol.
   2. Check the library for concentration and quality on a DNA Chip.
      NOTE: More precise and accurate quantification may be performed with sensitive fluorescent-based quantitation assays.
4. Library sequencing
   1. Normalize the samples to 2 nM and proceed to multiplexing according to the flow cell organization.
   2. Denature the samples at a concentration of 1 nM using 0.1 N NaOH for 5 min at RT.
   3. Dilute the samples at 10 pM. Load each sample on the flow cell at 10 pM.
   4. Proceed with sequencing with a high-throughput sequencing system that enables large-scale genomics. Here the run was an SRM run (SR: Single Read, PE: Paired-end Reads, M: multiplexed samples) of 51 cycles with 7 index bases read.
   5. Assess the quality of the sequencing with the external FastQC program¹³.
   6. After the trimming of adapter sequences, proceed with read alignments against a reference genome. Align against the eukaryotic cell genome to assess sample contamination and, if necessary, proceed with the trimming of the non-bacterial reads.
5. Statistical analysis
   1. Use several software programs available online to define sets of differentially expressed genes: R software, Bioconductor packages including DESeq2 and the PF2tools package (version 1.2.12) developed at the platform "Transcriptome et Epigénome" (Pasteur Institute, Paris).