DOI: 10.5061/dryad.hmgqnk9hz
title: Nipponaclerda biwakoensis infestation of Phragmites australis in the Mississippi River Delta, USA: Do fungal microbiomes play a role?

description: Recently, significant die-back of nonnative common reed, Phragmites
 australis, has been reported in the Mississippi River Delta (MRD),
 Louisiana, USA. This dieback has been attributed to an invasive scale
 insect, Nipponaclerda biwakoensis. We test whether fungi are involved in
 the recent infestation by this insect and subsequent die-offs of
 Phragmites australis. Several haplotypes of P. australis occur in the MRD,
 and the European (M) and Delta (M1) haplotypes appear to experience
 differing levels of N. biwakoensis infestation. We tested whether these
 haplotypes differed in their fungal microbiomes in both their leaf and
 stem tissues, and whether differences in fungal community composition were
 linked to the level of infestation using a metabarcoding Internal
 Transcribed Spacer (ITS) amplicon sequencing approach. Our analyses showed
 differences in fungal community composition and diversity between
 haplotypes and tissue types, but none of these differences were directly
 correlated with N. biwakoensis infestation severity. However, we did find
 that the European haplotype hosted higher putative pathogen loads in stem
 tissues compared to the Delta haplotype, which may confer resistance to
 herbivory, though it is possible that differences in infestation between
 haplotypes are due to morphology.
description: <b>Site and sample
 collection</b> On September 17,
 2018, shortly after peak biomass, we traveled to the Passe-a-Loutre
 Wildlife Management Area of the Mississippi River, Louisiana, USA.
 Passe-a-Loutre is a coastal freshwater marsh with a mean salinity of less
 than 1 ppt. <i>P. australis</i> is the dominant vegetation in
 the region (Knight et al., 2020). 20 samples of each haplotype (European
 and Delta) were collected at two separate sites (Table 1) for a total of
 80 individuals. The paired-haplotype plots at each site were identified by
 researchers at Louisiana State University on May 31, 2017 (Knight et al.,
 2020). Individuals of the European haplotype and Delta haplotype are
 easily differentiated in the MRD due to morphological and phenological
 differences, with the European haplotype exhibiting a much shorter, more
 gracile stature, earlier flowering time (all 40 European haplotype
 individuals collected were in flower at the time of sampling), and sparse
 or absent ligule hairs. The Delta haplotype is much taller,
 thicker-stemmed and possess dense ligule hairs (Hauber et al., 2011). No
 Delta haplotype individuals were in flower at the time of sampling. Culms
 of each haplotype were cut at the surface of the water if the plot was
 flooded (because scale insects do not occur under water) or at the soil in
 dry plots and placed individually in bags on wet ice for transport. Mean
 water depth at Site 1 was 19.8 cm and 2.6 cm at Site 2. Information on
 stand density and number of dead and living culms was determined for each
 individual using a circular quadrat which covers an area of ¼
 m<sup>2</sup> placed with the selected culm in the center. On
 the day of collection, samples were placed in a 4°C refrigerator for
 holding and small subsamples of internodal (hereafter, “stem”) and leaf
 tissue from each of the 80 individual culms collected were placed in a
 -80°C freezer for metabarcoding analysis. Using the
 remaining portions of the individuals, the number of <i>N.
 biwakoensis</i> insects per plant was determined by counting
 individuals every third internode and extrapolating to the entire culm.
 This was done by finding the average number of scales per node and
 multiplying by the number of internodes on the culm. Because the internode
 lengths differ within one individual, as well as between haplotypes, this
 value was standardized by the overall above-water height of the culm to
 determine scale density per centimeter of height. While the native scale
 insect <i>Aclerda holci</i> also occurs on <i>P.
 australis</i> in Louisiana, <i>N. biwakoensis</i> can be
 identified by its uniformly sclerotized and rounded abdomen (Knight et al.
 2018) and was the only scale species present on the collected <i>P.
 australis </i>samples. <b>Surface
 sterilization</b> For metabarcoding, 10 culms from
 each plot were selected by arranging all collected culms within that plot
 in order of lowest scale density to highest. Starting with the lowest,
 every-other individual was chosen for analysis. This ensured we had a
 range of infestation represented for each plot and each haplotype. A 10cm
 section of healthy leaf tissue and a 4cm internodal section of stem from
 each sample were selected and placed in a tea strainer. Plant tissue was
 surface sterilized following protocol developed for <i>Spartina
 patens</i> (Lumibao et al., 2018).<i> </i>Tissues were
 sequentially soaked in 95% ethanol for 10 seconds, 0.5% sodium
 hypochlorite solution for 2 minutes, 70% ethanol for 2 minutes, and rinsed
 in sterilized deionized water for 2 minutes. Samples were then dried on UV
 sterilized KIMTECH Kimwipes before placing in gamma-sterilized cryovials
 for storage at -80°C. <b>DNA
 extraction</b> Following surface sterilization,
 samples were homogenized using liquid nitrogen and a mortar and pestle
 sterilized between samples. Genomic DNA extractions for the leaf samples
 were done following Qiagen DNeasy<sup> ®</sup>
 PowerPlant<sup>®</sup> Pro Kit protocol using 50mg of tissue,
 Phenolic Separation Solution and 250µl of Solution IR. Leaf samples were
 eluted using 100µl of Solution EB. Due to low yields for stem tissue, 75mg
 of stem tissue was used and elution was done using 50µl of Solution EB.
 Extraction concentrations were determined using a ThermoFisher Scientific
 Qubit fluorometer and all samples above a concentration of 10ng/µl were
 standardized to 10ng/µl by mixing extraction product with additional
 Solution EB. <b>PCR</b>
 To allow
 for maximum flexibility in choosing region-specific primers and
 dual-indexing barcode combinations, sequencing libraries were created in a
 two-step process following U’ren and Arnold (2017). PCR1, or amplification
 and primer ligation, was done in duplicate with an annealing temperature
 of 54.0° C. The primer sequences used were ITS1F 5’ – CTT GGT CAT TTA GAG
 GAA GTA – 3’ and ITS2R 5’ – GCT GCG TTC TTC ATC GAT GC – 3’ (Gardes
 & Bruns, 1993; White et al., 1990). Sample duplicates were then
 pooled and dual indexing primers were added so that no two samples
 contained the same combination of indexing barcodes. Five nanograms of DNA
 from each sample were pooled into a common library along with four
 negative controls which were included for identification of possible
 contaminant sequences. The pooled library was then purified and
 concentrated using Agencourt AMpure XP Beads with Dynamag-2 Magnet and
 sent to Duke University for Illumina MiSeq v3 sequencing (300 bp paired
 reads). <b>Sequence
 analysis</b> <b>           
 </b>Amplicon sequence variants (ASV) were identified following the
 DADA2 ITS pipeline (version 1.8) (Callahan et al., 2016) in R (R Core
 Team, 2019). ASVs are “phylotypes” that are a single DNA sequence (not
 clustered by a sequence similarity threshold like operational taxonomic
 units, OTUs). Primers and adapters were trimmed using the Biostrings
 package in R (Pagès et al., 2020) and data was denoised and paired reads
 joined using DADA2. No further sequence trimming was done, because DADA2
 is robust to low-quality sequences through the incorporation of read
 quality information into its error model. Seven possible contaminant
 sequences were identified using the negative controls and removed using
 the decontam package in R (Davis et al., 2018). One sample was dropped due
 to very low reads. Taxonomy was assigned using UNITE 7.2 with a minimum
 bootstrap confidence of 70% (Nilsson et al., 2019) and functional guild
 assignments were obtained using FUNGuild (Nguyen et al., 2016).