Differential Transmission of Two Isolates of Wheat streak mosaic virus by FiveWheat Curl Mite Populations

Wosula, E. N., McMechan, A. J., Oliveira-Hofman, C., Wegulo, S. N., and Hein, G. L. 2016. Differential transmission of two isolates ofWheat streak mosaic virus by five wheat curl mite populations. Plant Dis. 100:154-158. Wheat streakmosaic virus (WSMV), typemember of the genusTritimovirus in the family Potyviridae, is an economically important virus causing annual average yield losses of approximately 2 to 3% in winter wheat across the Great Plains. The wheat curl mite (WCM), Aceria tosichella, transmits WSMV along with two other viruses found throughout the Great Plains of the United States. Two common genotypes of WSMV (Sidney 81 and Type) in the United States share 97.6% nucleotide sequence identity but their transmission relationships with the WCM are unknown. The objective of this study was to determine transmission of these two isolates of WSMV by five WCM populations (‘Nebraska’, ‘Montana’, ‘South Dakota’, ‘Type 1’, and ‘Type 2’). Nonviruliferousmites from each population were reared on wheat source plants mechanically inoculated with either Sidney 81 or Type WSMV isolates. For each source plant, individual mites were transferred to 10 separate test plants and virus transmission was determined by a double-antibody sandwich enzyme-linked immunosorbent assay. Source plants were replicated nine times for each treatment (90 individual mite transfers). Results indicate that three mite populations transmitted Sidney 81 at higher rates compared with Type. Twomite populations (Nebraska and Type 2) transmitted Sidney 81 and Type at higher rates compared with the other three populations. Results from this study demonstrate that interactions between virus isolates and mite populations influence the epidemiology of WSMV. Wheat streak mosaic virus (WSMV) is the type species of the genus Tritimovirus within the family Potyviridae, the largest group of plant viruses (Brunt et al. 1996; Stenger et al. 1998). This virus is a pathogen of wheat and other cereals in the Americas, Europe, Asia, and North Africa (Brunt et al. 1996; Dwyer et al. 2007; SánchezSánchez et al. 2001; Stenger et al. 1998). It is an especially serious pathogen in the Great Plains of North America, where it causes 2 to 3% annual yield loss in wheat (Appel et al. 2012). In serious outbreaks, WSMV often causes up to 100% yield loss in individual fields (Wegulo et al. 2008). The wheat curl mite (WCM), Aceria tosichella Keifer, is the only known vector of WSMV (Slykhuis 1955; Staples and Allington 1956). This mite also vectors two other viruses in wheat in the Great Plains: Wheat mosaic virus (WMoV), also known as High Plains virus, a tentative member of the genus Emaravirus (Seifers et al. 1997; Tatineni et al. 2014); and Triticum mosaic virus (TriMV; Poacevirus, Potyviridae) (Seifers et al. 2009; Tatineni et al. 2009). These three wheat viruses are widely spread across the Great Plains; however, WSMV has been shown to be the predominant virus in this complex (Burrows et al. 2008; Byamukama et al. 2013). Field populations of WSMV are complex and consist of numerous genotypes (Fuentes-Bueno et al. 2011; McNeil et al. 1996; Montana et al. 1996; Robinson and Murray 2013) but different genotypes rarely occurred (approximately 2%) in the same plant (McNeil et al. 1996). Three WSMV isolates (Sidney 81, Type, and El Batán 3) have been completely sequenced (Choi et al. 2001; Stenger et al. 1998). Sidney 81, the most characterized isolate (Brakke et al. 1990; Choi et al. 1999; Hall et al. 2001b; Stenger et al. 1998), was isolated in 1981 from an infected wheat plant from western Nebraska (Brakke et al. 1990). The Type isolate was originally isolated in 1937 from infected wheat plants fromKansas (McKinney 1937). Sidney 81 andType are the representative American isolates of WSMV, sharing 97.6% nucleotide sequence identity, and they produce similar symptoms in wheat (Choi et al. 2001; Hall et al. 2001a). Sidney 81 systemically infects and produces symptoms in Zea mays inbred line SDp2 but Type does not (Tatineni et al. 2011). El Batán, from Mexico (Sánchez-Sánchez et al. 2001), has diverged from the American strains and retains 79% nucleotide sequence identity to Sidney 81 and Type (Choi et al. 2001). According to McNeil et al. (1996), predominant WSMV isolates in Nebraska were indistinguishable from Sidney 81 but no isolates were identical to Type. A. tosichella is a complex of multiple cryptic lineages with diverse but distinct host ranges (Skoracka et al. 2012, 2014). In North America, two distinct genotypes of WCM have been identified and designated as ‘type 1’ and ‘type 2’ (Hein et al. 2012). The original designation of these two types was made from Australian mite populations by Carew et al. (2009) and later shown to match populations found in North America (Hein et al. 2012). These two genotypes are known to coexist as mixed populations within wheat fields and even within wheat heads (Schiffer et al. 2009; Siriwetwiwat 2006). These WCM genotypes vary in their response to different resistant genes in wheat (Harvey et al. 1999). They also differ in their ability to transmit the three wheat viruses found in the Great Plains. The type 2 genotype transmits TriMV and WMoV more efficiently compared with the type 1 genotype (McMechan et al. 2014; Oliveira-Hofman et al. 2015; Seifers et al. 2002). Oliveira-Hofman et al. (2015) found that transmission of WSMV by type 2 mites is higher from singly infected source plants than those coinfected with TriMV. In addition, Schiffer et al. (2009) demonstrated that the type 1 genotype from Australia was unable to transmit an unknown strain of WSMV. Available literature documents WSMV transmission using multiple mites (Seifers et al. 2002) or from source plants coinfected with WSMV or TriMV (Oliveira-Hofman et al. 2015). Little has been done to document specific effects of various virus strains or mite genotypes on WSMV transmission. In addition, much of this transmission work in recent years has been done on the same mite colonies that were maintained since 1996, according to Harvey et al. (1999). This transmission work has been of considerable value because it provided excellent comparisons of transmission of an established virus and two relatively new viruses. However, these colonies were collected almost two decades ago, and it is important to compare the old colonies to recently collected ones to determine whether there are any shifts in WSMV transmission rates. Corresponding author: E. N. Wosula; E-mail: [email protected] Accepted for publication 6 June 2015. http://dx.doi.org/10.1094/PDIS-03-15-0342-RE © 2016 The American Phytopathological Society 154 Plant Disease /Vol. 100 No. 1 Evaluating the impact of genetic variability for bothWSMV isolates and various WCM populations is essential to understand the impact of mite and virus variability on the epidemiology andmanagement of this virus–mite complex. The objective of this study was to determine transmission rates of two isolates of WSMV (Sidney 81 and Type) by five populations ofWCM, including three long-established populations (‘Montana’, ‘Nebraska’, and ‘South Dakota’) and two newly established populations (Type 1 and Type 2). Materials and Methods Mite populations.The studywas conducted as a randomized complete block design with a factorial arrangement of treatments consisting of five mite populations and two virus treatments (two WSMV virus strains). The five mite populations were designated Montana (MT), South Dakota (SD), Nebraska (NE), Type 1 (T1), and Type 2 (T2). MT, SD, and NE are populations that were established 19 years ago and are the same as those used to evaluate mite resistance in WCM-resistant wheat varieties (Harvey et al. 1999), transmission of WMoV (Seifers et al. 2002), mite genotype characterization (Hein et al. 2012), TriMV transmission (McMechan et al. 2014), and cotransmission of WSMV and TriMV (Oliveira-Hofman et al. 2015). The WSMV Sidney 81 strain was collected from western Nebraska in 1981 (Brakke et al. 1990). The Type isolate was originally isolated in 1937 from infected wheat plants in Kansas (McKinney 1937). T1 and T2 mites were established in the summer of 2011 by collecting 10 to 25 wheat tillers from a wheat field in each of three Nebraska counties (Box Butte, Scottsbluff, and Chase Counties). Field-collected wheat tillers were used to infest 14-day-old ‘Millennium’ wheat plants reared in 4-cm-diameter cone-tainers (Stuewe & Sons Inc., Tangent, OR) filled with standard greenhouse soil. Conetainers were covered with plastic cylindrical cages (5 cm in diameter and 50 cm in height), with two to three vents covered with Nitex screen (80-mm-mesh opening; BioQuip Products Inc., Compton, CA) after planting. One to two wheat tillers were used to infest each cone-tainer. Plants were transferred to a growth chamber with a photoperiod of 14 h of light and 10 h of darkness maintained at 27°C for 3 weeks. From this process, 13 different WCM colonies were established (1 in Box Butte County, 10 in Scottsbluff County, and 2 in Chase County). Subsequently, single mite transfers were done from these colonies onto 50 cone-tainers (approximately 4/colony) containing 14-day-old test plants. Each cone-tainer (clonal colony) was tested for WCM type using polymerase chain reaction (PCR) amplification and restriction enzyme digestion of an approximately 1,600-bp fragment of the nuclear ribosomal internal transcribed spacer (ITS) and associated 28S ribosomal DNA region. The PCR product was digested usingHhaI restriction enzyme (Promega Corp., Madison, WI), and restriction fragment length polymorphism (RFLP) scored by visual analysis was compared with the DNA ladder (Hein et al. 2012; Siriwetwiwat 2006). To establish nonviruliferous colonies, five eggs were transferred from each cone-tainer to a 14-day-old test plant. After 4 weeks, WCM of the same genotype were combined together onto 14-day-old plants in a single pot. Thereafter, mite colonies were maintained by transferring mites from the original pot to 14-day-old plants in new pots every 3 weeks. Plants from the original pots were tested for WSMV, TriMV, and WMoV using double-antibody sandwich enzyme-linked immunosorbent assay (DAS-ELISA) and reverse-transcription PCR, and all were confirmed negative for these viruses. The colonies were maintained on ‘Settler CL’ wheat sown in 15-cm-diameter plastic pots and isolated with cages. Each cage was assembled from plastic sheeting molded into a cylinder 15 cm in diameter and 60 cm high. Two 8-cm-diameter holes were cut on opposite sides of the cage, approximately one-third of the way up the cage. The top of the cage and the side vents were covered with Nitex screen. The five mite populations were reared in separate growth chambers held at 24 to 27°C, with a photoperiod of 14 h of light and 10 h of darkness and 30 to 40% relative humidity. Mite populations were maintained by transferring 50 mites per pot to 14-day-old wheat plants approximately every 3 weeks. Establishment of virus source plants. Settler CL wheat was sown in 4-cm-diameter cone-tainers filled with standard greenhouse soil prepared by mixing soil, sand, vermiculite, and peat moss in a ratio of 2:1:1:2. The cone-tainers were each seeded with three seeds, placed on greenhouse benches, and watered appropriately. The conetainers were covered with plastic cylindrical cages, as described above. The plants were fertilized three times aweekwith Scotts brand “Peter’s Professional”water-soluble 20-10-20 general-purpose fertilizer (Everris NA, Inc., Dublin, OH). After 10 days, wheat plants were thinned to one seedling per cone-tainer, and this plant was mechanically inoculated with WSMV Sidney 81 or Type isolate. Three plants per mite population and virus combination were used for run 1 but some source plants died or did not have enough mites; thereafter, five plants were used to ensure sufficient source plants and mites. Sidney 81 and Type were obtained from an infectious cDNA clone whose in-vitro-generated RNA transcripts were inoculated to wheat seedlings at the single-leaf stage (Choi et al. 1999; Tatineni et al. 2011). The virus inoculum was prepared by grinding infected wheat tissue in sterile distilled water (1:10 [wt/vol]) using a mortar and pestle. The plant leaves were lightly dusted with carborundum and inoculated by gently rubbing the inoculum onto leaves using the pestle. Four days after inoculation, 10 aviruliferous WCM were transferred from each of the five populations onto each of the source plants inoculated with WSMV Sidney 81 or Type strain. In all, 1 plant per treatment combination (10 plants per run) were also inoculated with sterile water and infested with mites to provide a mock as a check for contamination from virus or mites. To transfer mites to source plants, the mites were placed onto moist black insect-mounting triangles (1.3 by 0.4 mm) with a human eyelash attached to a wooden dowel. Triangles were placed into the axil of the newly emerging leaf of each source plant. After 24 h to allow mites to establish on the plants, plants were transferred to a growth chamber (14 h of light, 10 h of darkness, and 25 to 27°C). Mites were allowed to build up on source plants for a period of 4 weeks. Single-mite transfers. Test plants used in this study were 14-dayold wheat plants (twoto three-leaf stage) grown in cone-tainers (three seedlings per cone-tainer that were thinned to one at the time of mite transfers). This experiment was conducted three times (runs) for a total of 9 source plants and 90 test plants per treatment combination, except for NE mites and Sidney 81, SD mites and Type, T2 mites and Type, and T2 mites and Sidney 81, which each had a total of 8 source plants and 80 test plants due to either dead source plants or insufficient mite numbers in run 1. Large, active adult mites from each source plant were individually transferred to 10 separate test plants. To enable this extensive number of transfers to be accomplished in a reasonable amount of time, single-mite transfers were done by three different persons for each run (one replicate per person). After transfers were completed, source plants were placed individually into plastic Ziploc bags and stored at −20°C for subsequent WSMV assay via DAS-ELISA. Test plants were arranged in a randomized complete block design and held in a growth chamber (14 h of light and 10 h of darkness and 23 to 25°C). Mock plants were incorporated (one per five test plants) among the treatment plants to monitor potential contamination. The test plants were held in the growth chamber for 4 weeks before harvesting. They were cut at soil level and examined under a stereomicroscope to determine mite presence. Each test plant was put into individual Ziploc bags and stored at –20°C until virus testing by DAS-ELISA. Some one or two test plants in MT and Type, T1 and Type, T1 and Sidney 81, and T2 and Sidney 81 died and, therefore, were excluded from mite scoring or virus testing. Although plants were established in an insectand mite-free greenhouse and held in chambers with no other plants infested with either mites or viruses, source plants were also tested for TriMV and WMoV to ensure that they were not contaminated with these viruses. DAS-ELISA assays on test plants.Duplicate samples were tested for WSMV using DAS-ELISA. Positive WSMV controls consisted of wheat tissue inoculated with WSMV, and healthy wheat tissue was used as a negative control. ELISA plates (96-well Flat-Bottom ImmunoPlate;Maxisorp,Nunc, ThermoFisher Scientific Inc.,Waltham, Plant Disease / January 2016 155 MA) were coated with WSMV capture (primary) antibody (Agdia Inc., Elkhart, IN) in 1× carbonate buffer at 1:400 dilution and stored overnight at 4°C. Each sample was prepared by adding wheat tissue (approximately 0.15 g) along with general extraction buffer (Agdia Inc.) at a 1:10 (wt/vol) ratio to a mesh bag (Agdia Inc.). The sample was ground using a tissue homogenizer (Agdia Inc.). Plant tissue solution (100 ml) was added to each of two sample wells of the ELISA plate. WSMV alkaline phosphate conjugate (secondary) antibody (Agdia, Inc.) in general extraction buffer (1:400 dilution) was added (100 ml per well). Plates were incubated at 37°C for 1 h and rinsed seven times with phosphate-buffered saline-Tween buffer. Purine nucleoside phosphorylase (100 ml) was added to each well and incubated in the dark at room temperature for 1 h. Quantitative measurements of the reaction were determined using absorbance at 405 nm with a Multiscan FC Spectrophotometer (Thermo Fisher Scientific Inc.). Plants were considered positive for WSMV when the absorbance value was three times (or greater) that of a healthy (negative) control. Data analysis. Data analysis was performed using SAS software (version 9.4; SAS Institute Inc., Cary, NC). Proportions of WSMVinfected plants and presence of mites for the five mite populations and twoWSMV virus isolates were tested for differences using PROC GLIMMIX with binomial distribution. The LSMEANS statement was used to obtain least squares means and the Tukey-Kramer test at P = 0.05 was used for pairwise comparison of treatment means. Posthoc contrast analyses were also done to compare transmission rates among groups of mite populations. The effect of transfer person was analyzed as a fixed factor in a preliminary analysis to determine the appropriateness of considering these effects as replications within runs. In the final analysis, fixed factors were mite population and virus isolate, and run and replicate were included as random factors. Percent transmissionmeans and standard errors were obtained using the PROC MEANS statement.