Hybrid De Novo Genome Assembly for the Generation of Complete Genomes of Urinary Bacteria using Short- and Long-read Sequencing Technologies

Bacteria were cultured from urine collected from consenting women as part of institutional review board-approved studies 19MR0011 (UTD) and STU 032016-006 (UTSW).

1. Modified enhanced urine culture

NOTE: All culture steps must be carried out under sterile conditions. Sterilize all instruments, solutions, and media. Clean the work area with 70% ethanol, then set up a Bunsen burner and work carefully close to the flame to reduce the chances of contamination. Alternately, a class II biosafety cabinet may be used to maintain a sterile environment. Wear appropriate personal protective equipment (PPE) to avoid exposure to potentially pathogenic microbes.

1. Plating glycerol-stocked urine and colony isolation
   1. Thaw glycerol-stocked urine at room temperature (RT). Once thawed, vortex the sample for 5 s to mix. In sterile microcentrifuge tubes, prepare 1:3 and 1:30 dilutions of the urine in sterile 1x Phosphate-Buffered Saline (PBS) to a final volume of 100 µL.
      NOTE: Glycerol-stocked urine is prepared by mixing 500 µL of undiluted urine and 500 µL of 50% sterile glycerol in cryovials and storing at -80 °C.
   2. Pre-warm agar plates at 37 °C for 15 min before use. Please see Figure 1 for media types and culture conditions suitable to common urinary bacterial genera. Mix the diluted urine well by pipetting before plating, plate 100 µL of the diluted urine on the desired agar plate and spread the sample using sterile glass beads. Plate 100 µL of the 1x PBS diluent on a separate plate as a no growth control.
      NOTE: If attempting to culture common uropathogenic species (e.g., Escherichia coli, Klebsiella spp., Enterococcus faecalis, etc.), it is recommended to use chromogenic agar (Table of Materials) as it allows easy identification of uropathogenic bacterial species (Figure 1). Colistin Nalidixic Acid (CNA) or MRS agar are useful for isolating fastidious Gram-positive species (e.g., Lactobacillus spp.) from urine known to contain Gram-negative uropathogens, which may outcompete the fastidious species in non-selective agars.
   3. Incubate the plate inverted in the desired atmospheric condition at 35 °C for a period of 24 h for uropathogens and 3-5 days for fastidious bacteria (Figure 1).
   4. After the incubation period, remove the plates from the incubator. From each plate, pick the colonies that exhibit a unique color, morphology, or hemolytic patterns.
   5. Re-streak the bacterial colony using a sterile loop onto the corresponding agar and incubate the plate inverted for 2-5 days in the desired atmosphere to obtain well-isolated colonies.
      NOTE: If utilizing BAP for primary culture, patching colonies on chromogenic agar may provide useful information about the heterogeneity of the bacterial population in the sample.
2. Culturing in liquid broth and glycerol-stocking bacterial isolates
   1. Once the isolated colonies that match the morphology of the parent colony are obtained, pick a single colony and inoculate into 3 mL of liquid broth using a sterile inoculation loop. Refer to Figure 1 for broth capable of supporting the growth of common urinary microbiota genera. Seal the agar plates with parafilm and store them at 4 °C for 2-4 days. Incubate liquid cultures in the desired atmospheric conditions for 1-5 days until the culture is visibly turbid.
   2. After growth is observed, vortex the culture, and then add 1 mL of the overnight culture to 500 µL of sterile 50% glycerol in a 2 mL cryovial; seal and gently mix by inversion. Prepare two glycerol stocks for each colony (one serves as a backup) and store at -80 °C.

2. Identification of bacterial species by 16S rRNA gene Sanger sequencing

NOTE: Microbial identity can be alternatively confirmed using Matrix-Assisted Laser Desorption Ionization Time of Flight Mass Spectrometry (MALDI-TOF)²⁰.

1. Colony-Polymerase Chain Reaction (PCR)
   1. Prepare a 25 µL of the PCR reaction in PCR tubes by adding 12.5 µL of 2x Taq Polymerase Master Mix, 0.5 µL of 10 µM 8F primer, 0.5 µL of 10 µM 1492R primer (Table of Materials), and 11.5 µL of nuclease-free water²¹.
      NOTE: If performing PCR for multiple samples, make a reaction master mix of Taq Polymerase mix, primers, and sterile nuclease-free water. Then aliquot 25 µL into each PCR tube.
   2. To perform colony-PCR, swipe a well-isolated colony from the re-streak using a sterile toothpick or pipette tip. Resuspend the colony in the PCR reaction mix prepared in step 2.1.1. Gently mix. Collect the liquid at the bottom of the tube by a quick spin at 2000 x g.
      NOTE: Ensure the sample is free of air bubbles. Include a no-template control (NTC) sample containing the PCR reaction mix alone.
   3. Place the sample tubes in the thermocycler and run the following program: 95 °C for 3 min; 40 cycles of: 95 °C for 30 s, 51 °C for 30 s, and 72 °C for 1 min 30 s; 72 °C for 10 min; hold at 10 °C.
2. Gel extraction and species identification
   1. Upon completion of the PCR run, check the PCR product on a 1% agarose gel prepared in 0.5x Tris-Borate-EDTA (TBE) buffer. Prior to casting the gel, add ethidium bromide (EtBr). Then, cast the gel using combs for wells that hold at least 20 µL sample volume.
      CAUTION: EtBr is an intercalating agent suspected to be carcinogenic. Always wear gloves and PPE when handling it and dispose of materials containing EtBr according to the institution's guidelines.
   2. When the gel is set, place the gel in the electrophoresis tank filled with 0.5x TBE buffer and remove the comb. Load the 1 kb ladder in the first well and 10-20 µL of the PCR reaction into subsequent wells. Run at 100-140 V until resolved. Visualize the gel under UV light and confirm the presence of a clearly defined band at ~1.5 kb that is absent in the NTC well.
      CAUTION: UV rays are harmful to skin and eyes, use an appropriate guard when visualizing the gel and wear appropriate PPE.
      NOTE: Colony PCR may be unsuccessful for some bacteria; proceeding with PCR from isolated gDNA is an alternate option²².
   3. Excise the ~1.5 kb bands using a razor and transfer the gel cuttings into clean microcentrifuge tubes. Proceed with gel extraction protocol as per the manufacturer's instructions (Table of Materials). Measure the concentration of the purified DNA by microvolume spectrophotometer.
      NOTE: A concentration >10 ng/µL is desirable, and A260/280 between 1.7-2.0 is acceptable.
   4. Prepare two Sanger sequencing reactions for each sample, one using the 8F and the other using the 1492R primer in nuclease-free water according to the guidelines of any chosen Sanger sequencing service.
   5. Once the sequencing data is received, upload the DNA sequences to the NCBI Basic Local Alignment Search Tool (BLAST) website (blast.ncbi.nlm.nih.gov/Blast.cgi), choose Nucleotide BLAST (blastn), select the rRNA/ITS database 16S ribosomal RNA sequences (Bacteria and Archaea), and run the Megablast program. The isolate may be identified by the highest quality hit to a reference from the database.
      ​NOTE: Some bacterial species exhibit high identity in their 16S rRNA sequences and may be indistinguishable by this method alone. Speciation will require DNA homology and biochemical analyses to confidently distinguish members of the same genus²³.

3. Extraction of genomic DNA (gDNA)

NOTE: This section utilizes reagents and spin-columns provided in the gDNA extraction kit referenced in the Table of Materials for the high yield extraction of quality genomic DNA from diverse bacterial species. Provided below are recommended modifications and instructions.

1. Prepare kit reagents per manufacturer's instructions.
2. Prepare 3-10 mL cultures in appropriate sterile broth (Figure 1) by inoculating bacteria from well-isolated colonies into the media and incubating at the temperature and atmospheric pressure noted in Figure 1 until sufficient growth is observed.
3. After incubation, measure the optical density at 600 nm (OD₆₀₀) of the culture using a spectrophotometer²⁴.
   1. Prepare the sample for quantification by diluting overnight cultures in 1:10 ratio. Include a blank of the sterile culture media for measurement as well. Calculate the optical density by subtracting the blank reading from the sample reading and multiplying by the dilution factor of ten.
4. Using the OD₆₀₀ measurement and a pre-established OD₆₀₀ to CFU/mL ratio for the species, calculate how many milliliters of culture are necessary to obtain 2 x 10⁹ cells.
5. Centrifuge the required culture volume for 5 min at 5000 x g to pellet. Aspirate the supernatant and resuspend the pellet in 200 µL cold TE buffer (pre-chill on ice at the beginning of the procedure).
6. Centrifuge the sample for 2 min at 5000 x g. Remove the supernatant, and then resuspend the pellet in 180 µL of Enzymatic Lysis Buffer (ELB) and add 20 µL of pre-boiled RNase A (10 mg/mL). For efficient lysis of Gram-positive bacteria, add 18 µL of mutanolysin (25 kU/mL). Vortex well, and then incubate the samples at 37 °C on rotator for 2 h.
   NOTE: It is recommended to utilize the ELB described in the manufacturer's protocol for both Gram-positive and Gram-negative bacteria.
7. Proceed according to the manufacturer's instructions.
   NOTE: Repeat the elution steps for one or two more times to obtain additional gDNA yield, if desired.
8. Assess the quality of extracted gDNA as instructed in section 4 and store gDNA at 4 °C if it will be used within 1 week. Alternatively, keep gDNA at -20 °C for long-term storage.

4. Assessing the quality of extracted gDNA

1. To assess the quality by gel electrophoresis, prepare 1% agarose gel as described in subsection 2.2. Prepare the sample in a clean tube: mix 1-2 µL of extracted gDNA and 3 µL of 2x loading dye on parafilm. Run the gel once loaded, and then visualize it under UV light.
   NOTE: Successful gDNA extraction will be evident by a discrete band at the top of the gel and minimal smearing (Figure 2A). Smearing is indicative of shearing. If no gDNA band is evident and/or smearing is substantial, repeat gDNA extraction. Consider reducing incubation times in RNase A and Proteinase K. If two bands around 1.5-3 kb are observed, this suggests RNA contamination (Figure 2B). Prepare fresh RNase A and repeat extraction.
2. To assess the quality by microvolume spectrophotometer, measure gDNA concentration and absorbance ratio A260/280 by microvolume spectrophotometer. Concentrations >50 ng/µL and A260/280 between 1.7-2.0 are acceptable.
   NOTE: Low gDNA yield may be due to low input, high input, contamination of nucleases, insufficient lysis. Absorbance ratios above the range indicate RNA contamination. Repeat extraction if the gDNA quality is poor.
3. To assess the quality by fluorometer, follow the manufacturer's instructions to quantify gDNA concentration using High-Sensitivity assay kit and fluorometer instrument (Table of Materials). Concentration >50 ng/µL is desirable.

5. Paired-end next generation short-read sequencing and library preparation

NOTE: Short-read sequencing may be performed on various instruments at distinct read lengths and orientations. 150 bp (300 cycle) paired-end sequencing is recommended for bacterial WGS. Both library preparation and sequencing may be outsourced to core facilities or commercial laboratories.

1. Prepare sequencing library according to the manufacturer's instructions (Table of Materials). Follow the manufacturer's recommended final loading library concentration; however, a recommended modification is to load the pooled library at 1.8 pM for optimal read generation on NextSeq instruments.
2. Although optional, utilize a Bioanalyzer (Table of Materials) to assess the pooled library fragment distribution and ensure that the fragment size is 600 bp on average.

6. Nanopore MinION sequencing library preparation

1. Prepare the sequencing library according to the manufacturer's protocol (Table of Materials). Using two barcode expansion kits allows for multiplexing of up to 24 samples on a single flow cell. It is recommended to perform library preparation in two parts, 12 samples at a time when multiplexing 24 samples. All 24 samples may be pooled as described below.
   NOTE: Samples may be stored at 4 °C overnight upon finishing Native Barcode Ligation – this provides a stopping point in the protocol, if necessary. At the end of the Native barcode ligation section of the library preparation protocol, it is recommended to pool equimolar amounts of each sample up to the maximum DNA mass (ng) possible.
   1. To do so, quantify all the samples following barcode ligation using a fluorometer (Table of Materials) per manufacturer's instructions. Estimate the volume of the sample with the lowest dsDNA concentration, and then calculate the total dsDNA found in this sample. Use this number to determine the equimolar amounts of all the other samples that will be pooled together.
      NOTE: Because the equimolar calculation will maximize the amount of pooled dsDNA and thus yield a high-volume pool (>65 µL), cleanup is necessary to concentrate the pool.
2. dsDNA pool cleanup and concentration
   1. Add 2.5x volume of paramagnetic beads (Table of Materials) to the DNA pool, and then gently flick the tube to mix the contents. Place the tube in the rotator for 5 min at RT. Spin down the sample at 2000 x g and pellet on a magnet.
   2. Add 250 µL freshly prepared 70% ethanol (in nuclease free water), taking care not to disturb the pellet. Aspirate the ethanol and repeat the ethanol wash once.
   3. After the second aspiration, spin down the sample at 2000 x g and place it back on the magnet. Pipette off any residual ethanol and allow the sample to dry for approximately 30 s.
   4. Remove the tube from the magnet and resuspend the pellet in 60-70 µL of nuclease free water. Incubate at RT for 2 min. Pellet the sample on the magnet until the elute is clear, and then remove the elute and transfer into a clean 1.5 mL microcentrifuge tube.
   5. Quantify the concentrated pool using a fluorometer, and then prepare an aliquot to proceed to the adapter ligation step: prepare 700 ng of the sample in 65 µL final volume. Retain the remainder of the pool at 4 °C for a second run to be completed once the first run is finished.
   6. Proceed with adapter ligation as instructed by the manufacturer and load the sample on the flow cell. Start the sequencing run.
      NOTE: Aspirate air and ~200 µL of storage buffer from the flow cell priming port prior to the sample loading. This is critical for the successful flow cell priming and sample loading. Use a p1000 pipette and tips when drawing and depositing solutions through the priming port of the flow cell.
3. Sequence the library according to the manufacturer's instructions.
   1. Open the operating software for sequencing and click on Start. Input a name for the experiment, a recommended nomenclature includes the run date and user's name. Click on Continue to Kit Selection, select the appropriate library prep kit and barcode expansion pack(s) used, and then click on Continue to Run Options.
   2. Adjust the run length to 48 h if planning to prepare sufficient library for a second run (otherwise leave at default 72 h). Click on Continue to Basecalling.
   3. Check the basecalling option Config: Fast Basecalling and make sure that Barcoding is set to Enabled so that output FASTQ files will be trimmed of the barcode sequences and demultiplexed into separate directories based on barcode. Click on Continue to Output.
   4. Choose where to save output sequencing data. Expect approximately 30-50 Gb of data if only saving FASTQ output and >500 Gb of data if also saving FAST5 output. Uncheck the Filtering option Qscore: 7 | Readlength: Unfiltered if planning to proceed with filtering described in section 7.2, otherwise leave checked and adjust Readlength to 200.
   5. Click on Continue to Run Setup and review all the settings. If the settings are correct, click on Start, otherwise click on Back and make any necessary adjustments.
   6. If desired, the flow cell may be washed per the manufacturer's instructions and reloaded with the remaining pool. Repeat the steps in 6.2 for the remaining pool once the first run is complete and the flow cell has been washed.
      NOTE: When setting up the second run, adjust the Bias voltage to -250 mV per the manufacturer's recommendations for flow cells previously used in runs over 48 h.

7. Assessing and preparing reads

NOTE: A recommended directory structure is depicted in Figure 4. Create the directories found in the Desktop, namely, Long_Reads, Short_Reads and Trimmed_Reads, prior to proceeding with the computation steps below.

1. Short reads (Figure 3)
   NOTE: Short reads are generated in the FASTQ format. The files contain 4000 maximum reads per FASTQ. These are often zipped (.gz archive) and organized into multiple files. Depending on the platform, barcodes are typically trimmed. Some programs accept files in the zipped format, others may require their extraction prior to importing. The reads must pass quality control (QC) steps to ensure data accuracy during genome assembly. If CLC Genomics Workbench is not available, alternate programs may be used to trim and QC short reads such as Trimmomatic²⁵ or Trim Galore (https://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) for trimming and FastQC (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/) for evaluating read quality. Average short read coverage, estimated by multiplying number of reads by average read length and dividing by the genome size, is recommended to be >100x.
   1. Open Genomics Workbench software (Table of Materials) and import all paired-end short-read FASTQ files. Paired files will be generated automatically.
      1. Create a new folder under CLC_Data by clicking on the 신규 at the top toolbar and selecting Folder… to store the files. Name the folder as desired, a recommended convention is using the sample ID. Save all the output from the following steps to this folder.
      2. At the top toolbar, click on the Import button and select Illumina… Navigate to and select all short-read files that correspond to the sample. Make sure that the paired reads option is selected and uncheck the Remove failed reads option. Click on Next, select Save, and click on Next again. Choose to save the imported files in the new folder created in the previous step and click on Finish.
   2. Create a sequence list of all paired files for the isolate; this will concatenate read data into a single file for simplicity of analysis.
      1. At the top toolbar, click on the 신규 button and select Sequence List… On the directory list on the left, select the files to be concatenated and use the arrows to move those into the selected files list on the right. Click on Next, select Save, and click on Next again. Choose to save the sequence list and click on Finish.
      2. Once the sequence list is generated, immediately rename it with the sample ID.
   3. Run the QC for Sequencing Reads tool on the sequence list: This procedure will assess the overall quality parameters of the reads generated by short-read NGS.
      1. Search for the QC for Sequencing Reads tool in the toolbox menu (bottom-left window). Double-click on the tool, and then choose the sequence list to be analyzed and click on Next.
      2. Ensure all the output options are checked and choose Save under Results Handling. Click on Next and specify to save the output files, and then click on Finish.
   4. Run the Trim Reads tool on the sequence list: Trimming will be done based on quality, length, and ambiguity. This process assumes the barcodes used in sequencing have been trimmed prior to this step.
      1. Search for the Trim Reads tool in the toolbox (bottom-left window). Double click on Trim Reads, and then choose the sequence list to be analyzed and click on Next.
      2. Quality trimming: set the quality score limit to 0.01 and leave ambiguous nucleotides at 2. Click on Next.
         NOTE: Parameters may be adjusted at user discretion; these are the recommended settings.
      3. Uncheck Automatic Read-Through Adapter Trimming (only do this if adapters have been trimmed from the reads prior to import into CLC). Click on Next and check Discard Reads Below Length, use default 15.
      4. Click on Next, check Create Report, and then choose Save. Click on Next and specify where to save the output files. Click on Finish.
   5. Export the trimmed sequence list: subsequent hybrid assembly and analysis will be completed outside of CLC and requires trimmed short-read files to be exported.
      1. From the directory navigation on the top left, choose the trimmed file generated in step 7.1.4, and then click on Export at the top toolbar. Select Fastq for the export file type and click on Next. Check Export Paired Sequence List to Two Files. Then, click on Next and choose the Trimmed_Reads directory to export the files to. Click on Finish. Ensure that the trimmed short-read files were exported successfully as two files (R1 and R2) with the extension .fastq.
         NOTE: The trimmed sequence list must be exported into two files, typically designated by CLC as R1 and R2. This is critical as downstream hybrid assembly requires short-read data input to be set up as such.
      2. Rename the exported files, please refrain from the use of spaces and special characters in file names. For simplicity a recommended format is trimmed_short_file.R1.fastq.
2. Long (MinION) reads (Figure 3)
   NOTE: The following pipeline for the preparation of Long (MinION) sequencing reads for hybrid assembly utilizes NanoFilt and Nanostat programs²⁶ executed by the command-line. Install the tools prior to proceeding and be familiar with the basics of UNIX in order to execute these commands. Default terminals and Bash Shell are recommended. A lesson guide for common terminal commands and usage is found at Software Carpentry²⁷. The instructions below assume that the files generated will be named with the barcode nomenclature (NB01, NB02, etc.) and are saved in the Long_Reads directory. Alternatively, read filtering can be accomplished using MinKNOW when setting up the sequencing run. Average long read coverage is recommended to be >100x. Recommended average read length is >2000 bp; therefore, the number of long reads needed is lower than the number of short reads.
   1. Create new directories for each barcode used in the run (barcode01, barcode02, etc.) within the Long_Reads directory (Figure 4). Copy all of the .fastq files that correspond to each barcode into the appropriate folder. Combine all .fastq files for each barcode from every run.
   2. Open Terminal and navigate to the barcode directories within the Long_Reads directory using the cd command: cd Desktop/Long_Reads/barcode01
   3. Concatenate all .fastq files per barcode into a single .fastq file by executing the following command: cat *.fastq > NB01.fastq
      NOTE: This command combines all of the reads from each of the FASTQ files into one large, single FASTQ named NB01.fastq.
   4. Use NanoStat to assess read quality of the sample by executing the following command: NanoStat –fastq NB01.fastq
   5. Record the results by copying the output into a text or Word file for future reference.
   6. Use NanoFilt to filter MinION reads discarding reads with Q < 7 and length < 200 by executing the command: NanoFilt -q 7 -l 200 bp NB01.fastq | gzip > NB01 _trimmed.fastq.gz
   7. Run NanoStat on the trimmed file generated in step 7.2.6 by executing the command: NanoStat –fastq NB01 _trimmed.fastq.gz
   8. Record the results by copying the output into a text or Word file and compare to the results from step 7.2.4 to ensure that the filtering was successful (Table 1).
   9. Repeat steps 7.2.2 to 7.2.8 for each barcode used in the sequencing run.
      ​NOTE: The NB01_trimmed.fastq.gz file generated in step 7.2.6 will be used for hybrid assembly.

8. Generating hybrid genome assembly

NOTE: The following assembly pipeline utilizes Unicycler^19,28,29,30 to combine short and long reads prepared in sections 7.1 and 7.2 (Figure 3). Install Unicycler and its dependencies and execute the commands below. Short-read files exported in step 7.1.5 are assumed to be named trimmed_short_file.R1.fastq and trimmed_short_file.R2.fastq for simplicity.

1. Organize the short-read files and long-read files into a single directory named Trimmed_Reads. The directory must contain the following:
   1. A .fastq.gz file for trimmed long reads (generated in step 7.2.6).
   2. Two .fastq files (R1 and R2) for trimmed short reads (generated in step 7.1.5).
2. Navigate to the directory Trimmed_Reads that stores the read files using the cd command in Terminal: cd Desktop/Trimmed_Reads
   1. Once in the correct directory, zip the two short read files so they are also in the .fastq.gz format by executing the following command: gzip trimmed_short_file.R1.fastq
3. Repeat step 8.2 for both R1 and R2. Check that all the read files are now in the .fastq.gz format and verify that all the files match the same isolate.
4. Begin the hybrid assembly using Unicycler by running the following command:
   unicycler -1 trimmed_short_file.R1.fastq.gz -2 trimmed_short_file.R2.fastq.gz -l NB01 _trimmed.fastq.gz -o unicycler_output_directory
   ​NOTE: -o specifies the directory in which the Unicycler output will be saved, Unicycler will create this directory once the command is executed; do not generate the directory beforehand. Run time varies by computational power of the computer used as well as genome size and the number of reads. This may take anywhere from 4 h to 1 or 2 days. This protocol was performed on a CentOS Linux 7 machine with 250 Gb RAM, Intel Xeon (R) CPU with 2.5 GHz 12 practical cores and 48 virtual cores. Alternatively, personal computers with 16 Gb RAM and 2.6 GHz 6-core processors can compute these assemblies at a longer processing time.
5. When the run is complete, review the unicycler.log file to ensure no errors – record the number, size, and status (complete, incomplete) of the contigs generated.
   1. If incomplete contigs are identified (denoted as incomplete in the Unicycler log), re-run Unicycler in bold mode by adding the following flag to the command in step 8.4: –mode bold.
      ​NOTE: Bold mode will lower the quality threshold accepted for long read bridges during assembly; this may yield a complete assembly, but the assembly quality may be diminished. It is recommended to utilize bold mode only when necessary and as preliminary evidence for contig joining to be later confirmed by PCR.

9. Assessing assembly quality

NOTE: The following protocol utilizes Bandage³¹ and QUAST³², two programs that must be set up prior to use (Figure 2 and Figure 4). Bandage does not require installation once downloaded and QUAST requires familiarity with basic command-line usage. It is also recommended to assess genome completeness using Benchmarking Universal Single-Copy Orthologs (BUSCO)³³.

1. Bandage: Click on File. Then, choose Load Graph and select the assembly.gfa file that was saved to unicycler_output_directory generated by Unicycler in step 8.4. Once loaded, click on the Draw Graph button on the left-hand toolbar and look at how the contigs (called nodes) are connected and organized to evaluate if the assembly is complete (Figure 5).
   NOTE: Complete assemblies are represented by single circular contigs linked at both ends (Figure 5A,B). Incomplete assemblies have multiple contigs linked together or are linear (Figure 5C). Small linear contigs may not be incomplete as they may indicate linear extrachromosomal elements. Coverage, also called depth, will be noted in bandage and represents the relative abundance of the contigs to the chromosome, normalized in Unicycler to 1x.
2. QUAST
   1. Within the Terminal, navigate to the folder that stores the Unicycler output using the cd command: cd Desktop/Trimmed_Reads/unicycler_output_directory
      ​NOTE: Spaces are not permitted in the path to where the assembly is located, i.e., no directories leading to the Unicycler output can have spaces in their name. Alternatively, copy the assembly.fasta file to the Desktop for easy access.
   2. Run QUAST by executing the following command: quast assembly.fasta -o quast_output_directory
   3. Review the reports generated by QUAST in the output directory quast_output_directory.

10. Genome annotation

NOTE: The below annotation pipeline utilizes Prokka³⁴, a command-line tool that must be installed prior to usage. Alternatively, use Prokka through the automated GUI K-Base (Table of Materials) or annotate genomes via the web server RAST³⁵. If depositing genomes into NCBI, they will be automatically annotated using the Prokaryotic Genome Annotation Pipeline (PGAP)³⁶.

1. Navigate within the Terminal to the folder that stores the Unicycler output using the cd command (see step 9.2.1). Then, run Prokka by executing the following command: prokka —prefix sample_ID —outdir prokka_output_directory assembly.fasta
   NOTE: –prefix will name all output files based on the specified sample_ID. –outdir will create an output directory with the specified name where all Prokka output files will be saved; do not create an output directory for Prokka beforehand.
2. Review the annotations by opening the .tsv table and/or by uploading the .gff file generated into a sequence analysis software to visualize and analyze the annotations (Figure 6).
3. Specific types of annotations can be generated depending on genetic factors of interest. It is recommended to start with the user-friendly tools on the Center for Genomic Epidemiology (www.genomicepidemiology.org/) web server for preliminary analysis^{37,38,39,40,41}. Additional tools for detection of CRISPR-cas systems and prophage are available (Figure 3)^42,43.

11. Suggested practices for data democratization

1. When possible, deposit all the raw read data as well as assembled genomes in a public repository such as NCBI Sequence Read Archive (SRA) and Genbank. Genomes are automatically annotated via the PGAP pipeline during the NCBI deposition process.