Characterization and microsatellite marker development for Geosmithia obscura, a common bark and ambrosia beetle associate
Pietsch, Grace et al. (2022), Characterization and microsatellite marker development for Geosmithia obscura, a common bark and ambrosia beetle associate, Dryad, Dataset, https://doi.org/10.5061/dryad.nk98sf7w2
Background. Symbioses between Geosmithia fungi and wood-boring and bark beetles seldom result in disease induction within the plant host. Yet exceptions exist such as Geosmithia morbida, the causal agent of Thousand Cankers Disease (TCD) of walnuts and wingnuts and Geosmithia sp. 41, the causal agent of Foamy Bark Canker disease of oaks. Isolates of G. obscura were recovered from black walnut trees in eastern Tennessee and at least one isolate induced cankers following artificial inoculation. Due to the putative pathogenicity and lack of recovery of G. obscura from natural lesions, a molecular diagnostic screening tool was developed using microsatellite markers mined from the G. obscura genome.
Results. A total of 3,256 candidate microsatellite markers were identified (2236, 789, 137 di-, tri-, and tetra- motifs were identified, respectively), with 2011, 703, 101 di-, tri-, and tetra- motifs containing markers with primers. From these, 75 microsatellite markers were randomly selected, screened, and optimized, resulting in 28 polymorphic markers that yielded single, consistently recovered bands which were used in downstream analyses. Five of these microsatellite markers were found to be specific to G. obscura and did not cross-amplify into other, closely related species. Although the remaining tested markers could be useful, they cross-amplified within different Geosmithia species, making them not reliable for G. obscura detection.
Conclusion. Five novel microsatellite markers (GOBS9, GOBS10, GOBS41, GOBS43, GOBS50) were developed based on G. obscura genome. These species-specific microsatellite markers are available as a tool for use in molecular diagnostics and can assist future surveillance studies.
Genome sequencing, assembly, and microsatellite development
For whole-genome sequencing DNA from G. obscura isolate 6BE2, which originally was cultured from body wash samples from a X. crassiusculus beetle live-trapped in eastern Tennessee (Chahal et al. 2019), was extracted using Qiagen Blood and Cell Culture DNA Kit Maxi (Qiagen, Germantown, MD, USA), according to the protocol (Gazis et al., 2016). Libraries were prepared at the Michigan State University Genomics Core lab (https://rtsf.natsci.msu.edu/genomics/) using the Illumina TruSeq Nano DNA Library Preparation kit on a Perkin Elmer Sciclone G3 robot following the manufacturer’s recommendation. Completed libraries were checked for quality (QC) and quantified using a combination of Qubit dsDNA HS and Caliper LabChipGX HS DNA assays. All libraries were pooled in equimolar amounts based on QC and quantified using the Kapa Biosystems Illumina Library Quantification qPCR kit. Library sequencing was performed with Illumina HiSeq 4000 flow cell using a 2x150bp paired end format and a HiSeq 4000 SBS reagent kit. Base calling was completed using Illumina Real Time Analysis (RTA) v2.7.6 and output of RTA was demultiplexed and converted to FastQ format with Illumina Bcl2fastq v2.19.0.
The transcript quality of these reads was assessed using FastQC (Andrews, 2010) and error correction performed using default values with Bloom Filter Correction (BFC) (Li, 2015). Using the trimming program Skewer (Jiang et al., 2014) adapter sequences were removed and reads were filtered by requiring a minimal quality score of 20 in at least 70% of the bases. With the exception of minimal read length after trimming set to 30, all default parameters were used. Next, the transcripts were assembled using Assembly By Short Sequences (ABySS), specifically its paired-end option, abyss-pe, using a k-mer size of 81 and default settings for all other options (Simpson et al., 2009). Finally, sequences were masked for low complexity regions with Dustmasker (level of 1) (Morgulis et al., 2006).
Microsatellite markers were identified with a custom perl script (Staton and Ficklin, 2018) (Table 1). This script utilizes Primer3 (Rozen and Skaletsky, 2000) to search for di, tri, and tetra-repeating motifs, with primer product size range between 100-250 base pairs long (Untergasser et al., 2012). This script also produced text files containing the IDs and forward and reverse primers for the identified markers; these would be used to identify common regions between the different species’ genome scaffolds.
Fungal strain selection, DNA extraction, amplification and molecular confirmation
Following Gazis et al. (2018) protocol, axenic cultures from seven G. obscura isolates and 18 additional isolates of Geosmithia species (Table 2) were placed onto Difco™ Potato Dextrose Broth (PDB) (Becton, Dickinson and company, Sparks, MD, USA) at 22⁰C for up to two weeks, after which mycelium was harvested for DNA extraction. For species confirmation, GeneJet Genomic DNA Purification Kit (Thermo Fisher Scientific, Pittsburgh, PA, USA) was used, following manufacturer’s protocols with slight modifications. These modifications included increased proteinase K to 40 µL/sample and an extended overnight incubation period at 56°C. Samples were quantified using a nanodrop 1000 spectrophotometer (Thermo Fisher Scientific, Pittsburgh, PA, USA) and stored at -20⁰C until used. To confirm the identity of the Geosmithia isolates, the RNA operon was amplified and sequenced using the ITS primers ITS1F (Gardes and Bruns, 1993) and ITS4R (White et al., 1990), following Gazis et al. (2018) protocol. PCR product was visualized on a 2% agarose gel and sent to MCLAB (www.mclab.com) for cleaning and sequencing. Sequenced strands were assembled into contigs using Sequencher 5.0 (Gene Codes Corporation, Ann Arbor, MI, USA). Sequences were compared to the NCBI nucleotide database using BLAST search optimized to exclude uncultured/environmental sample sequences and to search sequences from type material. If species identity of 99-100% was not obtained, an unrestricted BLAST search was performed (Table 2). Additional Geosmithia spp. (G. obscura CBS121749, G. lavendula CBS344.49, G. pallida CBS260.33,) and other species (Penicillium [formerly Geosmithia] namyslowskii CBS686.85 and Talaromyces [formerly Geosmithia] viridulus CBS252.07) were acquired as DNA samples from The Dutch Centraalbureau voor Schimmelcultures (CBS) Fungal Biodiversity Centre collection or from previously verified DNA samples from our collection [G. obscura 14MCE1, G. sp. 23 4MN3, G. morbida GM182, G. morbida GM249, G. morbida GM250, and Rasamsonia argillacea (Stolk, H.C. Evans & T. Nilsson) Houbraken & Frisvad (formerly G. argillacea)].
Microsatellite characterization and cross-amplification
A total of 2815 microsatellite markers were identified with flanking primer sequences. Of those, 75 microsatellite markers (consisting of 25 di-, 25 tri-, and 25 tetra-nucleotide sequences) were randomly selected and screened to identify polymorphic markers. For the initial characterization, all primer pairs were tested using three G. obscura and one G. morbida isolates. PCR reactions were conducted using 4 µL GoTaq G2 Hot Start Colorless Master Mix (Promega Corporation, Madison, WI, USA), 1 µL each forward and reverse primers, 0.5 µL DMSO, 5 µL sterile water, and 1 µL genomic DNA providing a 12.5 µL sample volume. Samples were placed in a SimpliAmp ThermalCycler (Thermo Fisher Scientific) with the following protocol: 94°C for 3 min followed by 35 cycles of denaturation at 94°C for 40 sec, annealing at 55°C for 40 sec, and primer extension at 72°C for 30 sec, followed by 72°C for 4 min. PCR products were separated using a QIAxcel Capillary Electrophoresis System (Qiagen, Germantown, MD, USA) with a 25-500 bp size standard. Products with a relative fluorescence unit (RFU) of 100 or greater were scored as positive amplification. Only a subset of microsatellite markers (n=28) that were identified as polymorphic were further screened in the cross-amplification study. To accomplish this step, six G. obscura isolates along with 24 isolates from nine different Geosmithia species and three additional isolates outside Geosmithia were screened. Isolates were amplified using the PCR protocol described above and separated using QIAxcel Capillary Electrophoresis System with an RFU value of 100 or greater scored as positive. Number of alleles and haploid genetic diversity was obtained using the program GenAlEx 6.5 (Peakall and Smouse, 2012).
Abyss assembly of 9.1 million paired sequencing reads from DNA of G. obscura resulted in 5,752 unitigs spanning 28.9 Mb with an N50 of 24,134 and 47.4x coverage. The assembled sequences were screened for microsatellite development, from which 1,653 unitigs yielded at least one microsatellite marker, resulting in 3,256 candidate microsatellite markers. From this group, we identified 94 compound microsatellites, which were either located next to each other, or separated by less than 15 base pairs (bp), and 2,815 microsatellite markers with flanking primer sequences. Parameters for minimum number of replicates for each motif were established at 8 for dinucleotides, 7 for trinucleotides, and 6 for tetranucleotides. Using these baseline parameters, a total of 2236, 789, 137 di-, tri-, and tetra- motifs were identified respectively, with 2011, 703, 101 di-, tri-, and tetra- motifs containing markers with primers.
We tested 75 markers for amplification and the presence of polymorphic bands. All tested markers resulted in amplification, and a total of 36 markers were polymorphic (11 di-, 13 tri-, 12 tetra-nucleotides). Further optimization of the microsatellite markers yielded 28 markers with single, consistently recovered bands (Table 3), which were used to test cross-amplification of G. obscura markers into other Geosmithia species.
National Institute of Food and Agriculture, Award: Hatch project 1009630 (TEN00495)
Cooperative Agreement between the USDA FS Pacific Southwest Research Station and the University of Tennessee, Award: 15-CA-11272139-050
University of Tennessee Institute of Agriculture, Departments of Entomology and Plant Pathology and Plant Sciences