120

httpjournalstubitakgovtragriculture

Turkish Journal of Agriculture and Forestry Turk J Agric For(2016) 40 120-126copy TUumlBİTAKdoi103906tar-1410-113

New natural weed host Raphanus raphanistrum L (Brassicaceae) for Beet necrotic yellow vein virus and its vector Polymyxa betae Keskin

Nazlı Dide KUTLUK YILMAZ1 Emine KAYA ALTOP1 Colin James PHILLIPPO2 Huumlsrev MENNAN1

1Department of Plant Protection Faculty of Agriculture Ondokuz Mayıs University Samsun Turkey2Department of Horticulture Michigan State University East Lansing MI USA

Correspondence hmennanomuedutr

1 IntroductionBeet necrotic yellow vein virus (BNYVV) which is the agent of rhizomania disease is transmitted by Polymyxa betae Keskin and causes significant losses in sugar beet fields all over the world The disease is characterized in sugar beet plants by extensive rootlet proliferation from the main taproot and leaf chlorosis BNYVV is the type species of the genus Benyvirus and it contains four or five ssRNAs (Koenig et al 1986) Molecular analyses (restriction fragment length polymorphism single-strand conformation polymorphism and sequencing) of BNYVV isolates have revealed the existence of three strain groups (A B and P type) in Europe (Kruse et al 1994 Koenig et al 1995 Koenig and Lennefors 2000 Ward et al 2007) Rhizomania disease was first reported in sugar beet in Turkey in 1987 (Koch 1987) Since then it has spread to many major sugar beet-growing areas within the country especially the Western and Central Black Sea Region and the Central Anatolia and Marmara Regions

(Erdiller and Oumlzguumlr 1994 Kaya 2009 Kutluk Yilmaz and Arli Sokmen 2010) At present the disease is controlled by cultivating BNYVV-resistant sugar beet cultivars However resistance-breaking BNYVV strains have been found in cultivars carrying the Rz1 gene in the United States Europe (Liu and Lewellen 2007 Bornemann et al 2015) and Turkey (Kutluk Yilmaz et al 2012)

Both BNYVV and its vector P betae have limited host ranges A few members belonging to Amaranthaceae Asteraceae Caryophyllaceae Chenopodiaceae Portulacaceae Solanaceae and Poaceae were recorded as alternative hosts for BNYVV (Tamada and Baba 1973 Barr and Asher 1992 Hugo et al 1996 Legreve et al 2005 Mouhanna et al 2008) Until now BNYVV host studies on Brassicaceae such as Raphanus raphanistrum L Raphanus sativus L Brassica napus L and Sinapis arvensis L have failed to demonstrate infection by this virus or its vector (Barr and Asher 1992 Hugo et al 1996 Legreve et al 2005) R raphanistrum (wild radish) is a winter annual species

Abstract Rhizomania is an important virus disease of sugar beet (Beta vulgaris) caused by Beet necrotic yellow vein virus (BNYVV) The virus is transmitted to the roots of host plants by the plasmodiophorid Polymyxa betae During survey studies in September and October 2009 yellow vein banding symptoms were observed on wild radish (Raphanus raphanistrum) plants growing in spinach (Spinacia oleracea) fields with a history of rhizomania in Samsun Province in the Black Sea Region of Turkey To verify possible alternative hosts for BNYVV and P betae R raphanistrum and spinach plants and soil samples were collected BNYVV was detected in the leaf samples of field-collected R raphanistrum using the DAS-ELISA test This result was confirmed by RT-PCR and the Raphanus isolate was determined to be from the type A strain based on restriction fragment length polymorphism analysis The presence of P betae cystosori was not detected in the roots of R raphanistrum plants using a light microscopy technique but some objects resembling sporogenic plasmodia of P betae were observed in the infested root cells To confirm this result total RNA was extracted from the roots of these samples and tested by RT-PCR using Polymyxa-specific primers In contrast to the microscopy method all samples tested positive for P betae using RT-PCR Healthy seeds of R raphanistrum were planted in plastic pots containing soil from the same fields infested with BNYVV and the seedlings showed similar symptoms to those growing in natural conditions To our knowledge this is the first report of a BNYVV infection of R raphanistrum under natural conditions

Key words Rhizomania Polymyxa betae Brassicaceae Raphanus raphanistrum host

Received 28102014 AcceptedPublished Online 06082015 Final Version 01012016

Research Article

121

KUTLUK YILMAZ et al Turk J Agric For

belonging to Brassicaceae The broad ecological tolerance of these weeds such as quick germination rapid growth and abundant seed production make it a successful weed in many countries This weed a native of northern Europe and northern Asia is found in cultivated fields around the world (Cheam and Code 1995 Blackshaw 2001)

A major problem is that the virus particles survive inside resting spores of P betae for at least 15 years (Abe and Tamada 1986) In this long period some weed species act as alternative hosts for the virus and vector (Hugo et al 1996 Legreve et al 2005 Mouhanna et al 2008 Rysanek et al 2008) In 2009 several spinach fields with a history of rhizomania in Samsun Province in the Black Sea Region of Turkey exhibited yellow vein banding symptoms in wild radish and yellow-green mottling in spinach leaves The presence of these kinds of virus symptoms on plants in these fields raises the question of where BNYVV is present in wild radish and spinach crops Like sugar beet spinach (Spinacia oleracea L) also belongs to the family Chenopodiaceae In Italy BNYVV was identified in spinach plants in 1995 (Autonel et al 1995) and then the virus was found in spinach crops in California in 2010 (Liu et al 2010) More recently BNYVV was also detected in field-grown spinach in the western part of Turkey (Gumus et al 2014)

Improved control strategies for BNYVV require further knowledge of potential reservoirs of infection in sugar beet fields However reports on natural hosts of BNYVV based on large-scale surveys are scarce Maintenance of virus inoculum in weed species might provide a means for viruses to survive through the seasonal cycle and explain their prevalence during epidemics Therefore discovering the weed reservoirs of BNYVV may help us understand their epidemiology and provide basic information for implementation of effective management strategies Therefore the objective of this study was to test R raphanistrum for reservoirs of BNYVV and its vector P betae by using ELISA and RT-PCR tests

2 Materials and methods21 Plant and soil materialIn 2009 typical virus-like symptoms such as yellow-green mottling in spinach plants and yellow vein banding on R raphanistrum were observed in a spinach field in the Bafra district of Samsun Province in Turkey This area is well known from our previous studies to be highly contaminated by P betae and BNYVV For this reason a total of 25 R raphanistrum and 9 spinach plants were collected Soil samples were also taken from patches with plants displaying typical virus symptoms Additionally three soils infested with P betae and BNYVV and a noninfested soil (from the Ondokuz Mayıs University

Plant Protection Department collection) were used to grow sugar beets in a bait plant test as positive and negative controls in serological and molecular studies Samples were homogenized and used for bioassays under greenhouse conditions 22 Seed materialSeeds from Chenopodiaceae Brassicaceae Amaranthaceae and Portulacaceae were used in this study and supplied from Ondokuz Mayıs University Plant Protection Department collection23 Bait plant techniqueSeeds were sown separately into 500-mL plastic pots containing rhizomania-infested soil in three replications with ten seeds per plastic pot The pots were placed in a controlled glasshouse at 19 degC (night) and 25 degC (day) After 8 weeks plants were harvested and their roots were carefully washed in running tap water One section of the roots from each bait plant was checked for the presence of P betae cystosori by light microscope Another part was tested for the presence of BNYVV by ELISA Plants that tested positive for BNYVV or P betae were assessed again in a second test to determine if the virus and vector could be transmitted back to sugar beets For this purpose dried roots of infected plants were cut into small fragments and mixed with equal parts sterile sand and soil into which a BNYVV-susceptible sugar beet seed (Arosa) was sown After 8 weeks infections by P betae and BNYVV were assessed

Besides this sugar beet seeds of rhizomania-susceptible Arosa were planted into 300-mL pots of three BNYVV- and P betae-infested soils Noninfested soil samples mixed with sterile sand (11) were used as positive and negative controls in serological and molecular studies24 Serological tests DAS-ELISA was performed for BNYVV using polyclonal antiserum (Sediag France) according to Clark and Adams (1977) A sample was considered positive when the absorbance value at 405 nm was more than two times the mean of the negative controls (Meunier et al 2003) 25 Microscopy analysis of Polymyxa betaeRoot samples were stained with lactophenol containing 01 acid fuchsin and examined under a light microscope (Leica Switzerland) to detect the presence of P betae cystosori (Abe and Tamada 1986) 26 Total RNA isolationTotal RNA was extracted from the roots and leaves of R raphanistrum and the roots of sugar beet that had previously tested positive for BNYVV in ELISA Total RNA was extracted from 100 mg of frozen tissue using the RNeasy Mini Kit (QIAGEN USA) according to the manufacturerrsquos instructions

122

KUTLUK YILMAZ et al Turk J Agric For

27 RT-PCR for detection of BNYVV and Polymyxa betaeOne-step RT-PCR was performed as described in the manufacturerrsquos manual (QIAGEN) For the detection of BNYVV the primers proposed by Kruse et al (1994) were used (forward primer GTGATATATATGTGAGGACGCT and reverse primer CCGTGAAATCACGTGTAGTT) which amplified RNA-3 The RT-PCR reaction included the following 155 microL of RNase-free water 1 microL of dNTPs mix (400 microM) 5 microL of 5X Q solution 025 microL each of forward and reverse primers (06 microM) 1 microL of QIAGEN OneStep RT-PCR enzyme mix and 2 microL of RNA sample Reverse transcription consisted of 30 min at 50 degC and 15 min at 95 degC We then ran 35 cycles composed of denaturation for 30 s at 94 degC annealing for 30 s at 53 degC and elongation for 1 min at 72 degC A final elongation of 7 min at 72 degC was also performed

For the detection of Polymyxa in roots of R raphanistrum and sugar beet oligonucleotides flanking 58s-rDNA and internal transcribed spacers (ITS) were used This includes upstream 5`-GAGGCATGCTTCCGAGGGCTCT-3` and downstream primer 5`-CTGCGGAAGGAT-CATTAGCGTT-3`(Ward and Adams 1998) The RT-PCR reaction included the following 152 microL of RNase-free water 1 microL of dNTPs mix (400 microM) 5 microL of 5X Q solution 015 microL each of forward and reverse primers (06 microM) 1 microL of QIAGEN OneStep RT-PCR Enzyme mix and 25 microL of RNA sample Reaction conditions were the same as used for the amplification of RNA-3 except for annealing temperature (55 degC)

After PCR amplification 2 microL of loading buffer was added to the PCR products The samples were analyzed on 12 ethidium bromide agarose gel in 1X Tris-borate-EDTA buffer using the Gel Doc 2000 Systems (Bio-Rad USA) (Ward and Adams 1998)

28 Restriction fragment length polymorphism analysisTo produce a PCR product for restriction fragment length polymorphism (RFLP) analysis the primers proposed by Kruse et al (1994) were used to amplify the RNA-3 as described above which was then cut separately with enzymes EcoRI BamHI and MspI 3 Results and discussion31 Detection of BNYVVA total of 9 spinach plants and 25 R raphanistrum samples were collected from the spinach field and tested by DAS-ELISA In this study BNYVV was detected in 20 samples of R raphanistrum collected from the field (Figure 1A) On the other hand 7 of the field-grown spinach samples were infected with BNYVV Generally BNYVV-positive spinach plants have yellow-green mottling symptom on their leaves Although spinach is grown in many regions of Turkey BNYVV has only been recently recorded in spinach production areas in İzmir and Manisa provinces located in the western part of Turkey (Gumus et al 2014) In the ELISA tests with the R raphanistrum samples color development was very slow For this reason the DAS-ELISA test microtiter plates were read following 2 h substrate incubation at room temperature and then again following overnight substrate incubation at 4 degC Even after overnight incubations relatively low A405 values were obtained for most of the samples The positive controls supplied by the Sediag Biochemica and root samples from the sugar beet bait plants grown in BNYVV-infested soils gave satisfactory positive results at 2 h and 16 h substrate incubations The reason for the low ELISA values could be low virus titers in R raphanistrum samples In addition RT-PCR assay to detect BNYVV and P betae was carried out on total RNA extracted from leaves and root samples from R raphanistrum and root samples from the sugar beet bait All samples tested positive for BNYVV

A B

Figure 1 Yellow vein banding in field-grown (A) and greenhouse-grown (B) BNYVV-infected Raphanus raphanistrum

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KUTLUK YILMAZ et al Turk J Agric For

in this RT-PCR detection assay (Figure 2) Until now BNYVV host studies on Brassicaceae members such as R raphanistrum R sativus B napus and S arvensis have failed to demonstrate infection by this virus or its vector (Barr and Asher 1992 Hugo et al 1996 Legreve et al 2005) However BNYVV and P betae were both detected in Capsella bursa-pastoris (L) Medik and Thlaspi arvense L which belong to the family Brassicaceae in a bait plant test using RT-PCR by Legreve et al (2005) However in this study neither BNYVV nor its vector were found in these species by ELISA or light microscopy respectively (Legreve et al 2005) Later on Mouhanna et al (2008) found that C bursa-pastoris was infested with P betae and BNYVV using bait plant test and back transmission studies To our knowledge this is the first report of a BNYVV infection of R raphanistrum in the family Brassicaceae under natural conditions 32 RFLP analysis of BNYVVKruse et al (1994) showed that RNA-3 of B type strains is cut by BamHI EcoRI StyI and MspI In contrast the A type strains were not cut by any of these restriction enzymes Our BNYVV isolates were not digested by these restriction enzymes suggesting that the isolate from R raphanistrum belongs to the A type strain (Figure 3) The A type the most widespread strain (Schirmer et al 2005) has been previously detected in Turkey (Kruse et al 1994 Kutluk Yilmaz et al 2007) the B type strain is more restricted and mainly found in Germany France (Kruse et al 1994 Koenig et al 2008) Japan (Miyanishi et al 1999) and China (Li et al 2008)33 Detection of P betae in infested rootsRoots of R raphanistrum that were collected from a spinach field were inspected for the presence of P betae using light microscopy P betae cystosori were not detected in roots of any of these plants though some

objects resembling sporogenic plasmodia of P betae were described in the root cells of R raphanistrum (Figure 4) To confirm this result sugar beet bait plant and field-grown R raphanistrum samples were subjected to RT-PCR assay with Polymyxa-specific primers (Ward and Adams 1998) to determine whether they were infected with Polymyxa spp Ward and Adams (1998) emphasized that the products of amplification using these primers on isolates of Polymyxa graminis types I and II and P betae could all be distinguished from one another on 12 agarose gels Therefore a 250-bp DNA fragment expected for the ITS

M 1 2 3 4

750-bp

1000-bp

1500-bp

Figure 2 Confirmation of the presence of BNYVV in leaves of R raphanistrum using RT-PCR analysis M DNA size marker The arrow indicates the BNYVV-specific 1218-bp DNA fragment Lane 1 negative control (water) Lane 2 BNYVV-containing R raphanistrum leaves Lane 3 healthy control (noninfected sugar beet cultivar Arosa) Lane 4 positive control (sugar beet infected with BNYVV)

M 1 2 3 4 5 6 7 8

Figure 3 The results of RFLP analysis of the RT-PCR products of BNYVV RNA-3 M 1-kb ladder (Promega USA) The arrow indicates the BNYVV RNA-3-specific 1218-bp DNA fragment Lanes 1ndash4 RNA from roots of R raphanistrum (EcoR1 BamH1 Msp1 and uncut PCR product) Lanes 5ndash8 RNA from the roots of the bait sugar beet plant grown in BNYVV-infested soil (EcoR1 BamH1 Msp1 and uncut PCR product)

Figure 4 P betae sporogenic plasmodia in root cells of R raphanistrum

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KUTLUK YILMAZ et al Turk J Agric For

between the 58 s and the 18 s rRNA of P betae was obtained from the roots of the susceptible sugar beet cultivar grown in rhizomania-infested soils and R raphanistrum in this assay (Figure 5) Rysanek et al (2008) suggested that the PCR method would be useful for verifying the presence of P betae in the case of low numbers of cystosori in host range studies Moreover in some plant species plasmodia or zoosporangia could be present without cystosori PCR could reveal such hidden infections Similarly Legreve et al (2008) found that BNYVV and P betae were detected by RT-PCR in some species belonging to different families while the virus and vector went undetected by ELISA and microscopy respectively We suggest that viruliferous P betae zoospores were able to penetrate the roots of R raphanistrum plants and introduce the virus but the protozoa were unable to produce cystosori and complete their life cycle in the roots of wild radish An incomplete infection similar to our result was reported with P betae in Gomphrena globosa (Al Musa and Mink 1981) and P graminis in Arachis hypogaea (groundnut) (Thouvenel and Fauquet 1981) Later on Barr et al (1995) reported similar observations about early P betae developmental stages without cystosori in two wild beet species Beta patellaris and Beta procumbens Although the wild radish leaves became infected naturally with BNYVV in the field they were unable to provide sources of virus infection to other plants whereas comparable sugar beet roots were able to do so There are various factors such as environmental conditions the lack of vital nutritional

factors or insufficient nutrients which might have limited the development of pathogen structures (Barr et al 1995)34 Bait plant and back inoculation testsAmong all Brassicaceae species tested for P betae and BNYVV host status sugar beet spinach Chenopodium album L Chenopodium murale L Amaranthus retroflexus L and Portulaca oleracea L were added to the bait plant test The Table shows the species that became infected with either P betae or BNYVV In this study P betae cystosori were found by light microscopy only in sugar beet C album A retroflexus and P oleracea Interestingly we could not detect either P betae or BNYVV in spinach plants From all tested species infection by BNYVV was only detected in sugar beet and R raphanistrum using ELISA R raphanistrum plants also showed symptoms of yellow vein banding under controlled conditions similar to natural conditions (Figure 1B) However P betae was not detected using the light microscopy technique in these plant roots Neither BNYVV nor P betae was detected in the other tested Brassicaceae species R sativus E sativa B napus B rapa B juncea and B oleracea (Table)

Besides this we tried using roots of a plant infected with a vector or the virus (sugar beet C album A retroflexus P oleracea and R raphanistrum) to see if they had the potential to act as an alternative host for P betae and BNYVV to transmit to susceptible sugar beet plants Transmission back to sugar beet occurred only from B vulgaris (Table)

In rhizomania disease the virusndashvectorndashhost relationships are very complex Over the last decades the existence of different genotypic groups has become obvious and they correspond to differences in pathogenicity and specific geographical region BNYVV isolates fall into three strain groups A B and P types (Kruse et al 1994 Koenig et al 1995 Koenig and Lennefors 2000 Ward et al 2007) In the last years resistance-breaking BNYVV strains have been found in cultivars carrying the Rz1 gene in the United States Europe (Liu and Lewellen 2007 Bornemann et al 2015) and Turkey (Kutluk Yilmaz et al 2013) Beside this the performance of viruliferous P betae in soils is influenced by many different biotic and abiotic factors Gerik and Duffus (1988) described differences in vectoring abilities of P betae populations depending on their origin There are also subgroups of P betae with specific host ranges such as betae amaranthi and the family Portulacaceae (Barr 1979 Abe and Tamada 1986 Abe and Ui 1986) Moreover weeds are populations with a wide range of genetic variability In this respect this appears to be the first report of R raphanistrum from the family Brassicaceae acting as a host of BNYVV under natural conditions The Raphanus isolate was assigned to the A type strain based on RFLP analysis Although P betae cystosori were not found in the roots of

M 1 2 3 4 5

1000- bp

750- bp

500- bp

Figure 5 RT-PCR analysis for detection of P betae M 1-kb DNA ladder (Promega) The arrow indicates the P betae-specific 250-bp DNA fragment Lane 1 root RNA from R raphanistrum Lane 2 negative control (healthy sugar beet roots) Lanes 3ndash5 positive controls (BNYVV-containing sugar beet roots)

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KUTLUK YILMAZ et al Turk J Agric For

R raphanistrum it may infect wild radish roots without cystosori development Therefore R raphanistrum seems to be a relatively inefficient host for rhizomania compared to sugar beet Nonetheless knowing that this weed is a

host of BNYVV may help to understand its epidemiology and could provide basic information for implementation of effective management strategies Further work is needed in the search for natural hosts of BNYVV and P betae

Table ELISA and microscopy results for the bait plant and back inoculation to B vulgaris tests

Family Plant species Presence of P betae cystosori (microscopy)

Infection byBNYVV (ELISA)

Back inoculation B vulgaris

Chenopodiaceae Beta vulgaris L cv Arosa + + +

Chenopodiaceae Spinacia oleracea L - - nt

Chenopodiaceae Chenopodium album L + - -

Chenopodiaceae Chenopodium murale L - - nt

Amaranthaceae Amaranthus retroflexus L + - -

Portulacaceae Portulaca oleracea L + - -

Brassicaceae Raphanus raphanistrum L - + -

Brassicaceae Raphanus sativus L - - nt

Brassicaceae Brassica napus L (B-12 accession) - - nt

Brassicaceae Brassica napus (B-2 accession) - - nt

Brassicaceae Brassica juncea (L) Czern (B-4 accession) - - nt

Brassicaceae Brassica juncea (B-7 accession) - - nt

Brassicaceae Brassica juncea (B-5 accession) - - nt

Brassicaceae Brassica oleracea L - - nt

Brassicaceae Brassica rapa L - - nt

Brassicaceae Eruca sativa L - - nt

nt not tested

References

Abe H Tamada T (1986) Association of Beet necrotic yellow vein virus with isolates of Polymyxa betae Keskin Ann Phytopathol Soc Jpn 52 235ndash247

Abe H Ui T (1986) Host range of Polymyxa betae Keskin strains in rhizomania-infected soils of sugar beet fields in Japan Ann Phytopathol Soc Jpn 52 394ndash403

Al Musa AM Mink GI (1981) Beet necrotic yellow vein virus in North America Phytopathology 71 773ndash776

Autonel CR Turina M Fantino MG (1995) Spinach virus infections in Italy Inf Fitopatol 45 43ndash47

Barr DJ (1979) Morphology and host range of Polymyxa graminis Polymyxa betae and Ligniera pilorum from Ontario and some other areas Can J Plant Pathol 1 85ndash93

Barr KJ Asher MJC (1992) The host range of Polymyxa betae in Britain Plant Pathol 41 64ndash68

Barr KJ Asher MJC Lewis BG (1995) Resistance to Polymyxa betae in wild Beta species Plant Pathol 44 301ndash307

Blackshaw RE (2001) Influence of wild radish on yield and quality of canola weed Weed Sci 50 344ndash349

Bornemann K Hanse B Varrelmann M Stevens M (2015) Occurrence of resistance-breaking strains of Beet necrotic yellow vein virus in sugar beet in Northwestern Europe and identification of a new variant of the viral pathogenicity factor P25 Plant Pathol 64 25ndash34

Cheam AH Code GR (1995) The biology of Australian weeds 24 Raphanus raphanistrum L Plant Prot Q 10 2ndash13

Clark M Adams AM (1977) Characteristics of microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses J Gen Virol 34 475ndash483

121

KUTLUK YILMAZ et al Turk J Agric For

belonging to Brassicaceae The broad ecological tolerance of these weeds such as quick germination rapid growth and abundant seed production make it a successful weed in many countries This weed a native of northern Europe and northern Asia is found in cultivated fields around the world (Cheam and Code 1995 Blackshaw 2001)

A major problem is that the virus particles survive inside resting spores of P betae for at least 15 years (Abe and Tamada 1986) In this long period some weed species act as alternative hosts for the virus and vector (Hugo et al 1996 Legreve et al 2005 Mouhanna et al 2008 Rysanek et al 2008) In 2009 several spinach fields with a history of rhizomania in Samsun Province in the Black Sea Region of Turkey exhibited yellow vein banding symptoms in wild radish and yellow-green mottling in spinach leaves The presence of these kinds of virus symptoms on plants in these fields raises the question of where BNYVV is present in wild radish and spinach crops Like sugar beet spinach (Spinacia oleracea L) also belongs to the family Chenopodiaceae In Italy BNYVV was identified in spinach plants in 1995 (Autonel et al 1995) and then the virus was found in spinach crops in California in 2010 (Liu et al 2010) More recently BNYVV was also detected in field-grown spinach in the western part of Turkey (Gumus et al 2014)

Improved control strategies for BNYVV require further knowledge of potential reservoirs of infection in sugar beet fields However reports on natural hosts of BNYVV based on large-scale surveys are scarce Maintenance of virus inoculum in weed species might provide a means for viruses to survive through the seasonal cycle and explain their prevalence during epidemics Therefore discovering the weed reservoirs of BNYVV may help us understand their epidemiology and provide basic information for implementation of effective management strategies Therefore the objective of this study was to test R raphanistrum for reservoirs of BNYVV and its vector P betae by using ELISA and RT-PCR tests

2 Materials and methods21 Plant and soil materialIn 2009 typical virus-like symptoms such as yellow-green mottling in spinach plants and yellow vein banding on R raphanistrum were observed in a spinach field in the Bafra district of Samsun Province in Turkey This area is well known from our previous studies to be highly contaminated by P betae and BNYVV For this reason a total of 25 R raphanistrum and 9 spinach plants were collected Soil samples were also taken from patches with plants displaying typical virus symptoms Additionally three soils infested with P betae and BNYVV and a noninfested soil (from the Ondokuz Mayıs University

Plant Protection Department collection) were used to grow sugar beets in a bait plant test as positive and negative controls in serological and molecular studies Samples were homogenized and used for bioassays under greenhouse conditions 22 Seed materialSeeds from Chenopodiaceae Brassicaceae Amaranthaceae and Portulacaceae were used in this study and supplied from Ondokuz Mayıs University Plant Protection Department collection23 Bait plant techniqueSeeds were sown separately into 500-mL plastic pots containing rhizomania-infested soil in three replications with ten seeds per plastic pot The pots were placed in a controlled glasshouse at 19 degC (night) and 25 degC (day) After 8 weeks plants were harvested and their roots were carefully washed in running tap water One section of the roots from each bait plant was checked for the presence of P betae cystosori by light microscope Another part was tested for the presence of BNYVV by ELISA Plants that tested positive for BNYVV or P betae were assessed again in a second test to determine if the virus and vector could be transmitted back to sugar beets For this purpose dried roots of infected plants were cut into small fragments and mixed with equal parts sterile sand and soil into which a BNYVV-susceptible sugar beet seed (Arosa) was sown After 8 weeks infections by P betae and BNYVV were assessed

Besides this sugar beet seeds of rhizomania-susceptible Arosa were planted into 300-mL pots of three BNYVV- and P betae-infested soils Noninfested soil samples mixed with sterile sand (11) were used as positive and negative controls in serological and molecular studies24 Serological tests DAS-ELISA was performed for BNYVV using polyclonal antiserum (Sediag France) according to Clark and Adams (1977) A sample was considered positive when the absorbance value at 405 nm was more than two times the mean of the negative controls (Meunier et al 2003) 25 Microscopy analysis of Polymyxa betaeRoot samples were stained with lactophenol containing 01 acid fuchsin and examined under a light microscope (Leica Switzerland) to detect the presence of P betae cystosori (Abe and Tamada 1986) 26 Total RNA isolationTotal RNA was extracted from the roots and leaves of R raphanistrum and the roots of sugar beet that had previously tested positive for BNYVV in ELISA Total RNA was extracted from 100 mg of frozen tissue using the RNeasy Mini Kit (QIAGEN USA) according to the manufacturerrsquos instructions

122

KUTLUK YILMAZ et al Turk J Agric For

27 RT-PCR for detection of BNYVV and Polymyxa betaeOne-step RT-PCR was performed as described in the manufacturerrsquos manual (QIAGEN) For the detection of BNYVV the primers proposed by Kruse et al (1994) were used (forward primer GTGATATATATGTGAGGACGCT and reverse primer CCGTGAAATCACGTGTAGTT) which amplified RNA-3 The RT-PCR reaction included the following 155 microL of RNase-free water 1 microL of dNTPs mix (400 microM) 5 microL of 5X Q solution 025 microL each of forward and reverse primers (06 microM) 1 microL of QIAGEN OneStep RT-PCR enzyme mix and 2 microL of RNA sample Reverse transcription consisted of 30 min at 50 degC and 15 min at 95 degC We then ran 35 cycles composed of denaturation for 30 s at 94 degC annealing for 30 s at 53 degC and elongation for 1 min at 72 degC A final elongation of 7 min at 72 degC was also performed

For the detection of Polymyxa in roots of R raphanistrum and sugar beet oligonucleotides flanking 58s-rDNA and internal transcribed spacers (ITS) were used This includes upstream 5`-GAGGCATGCTTCCGAGGGCTCT-3` and downstream primer 5`-CTGCGGAAGGAT-CATTAGCGTT-3`(Ward and Adams 1998) The RT-PCR reaction included the following 152 microL of RNase-free water 1 microL of dNTPs mix (400 microM) 5 microL of 5X Q solution 015 microL each of forward and reverse primers (06 microM) 1 microL of QIAGEN OneStep RT-PCR Enzyme mix and 25 microL of RNA sample Reaction conditions were the same as used for the amplification of RNA-3 except for annealing temperature (55 degC)

After PCR amplification 2 microL of loading buffer was added to the PCR products The samples were analyzed on 12 ethidium bromide agarose gel in 1X Tris-borate-EDTA buffer using the Gel Doc 2000 Systems (Bio-Rad USA) (Ward and Adams 1998)

28 Restriction fragment length polymorphism analysisTo produce a PCR product for restriction fragment length polymorphism (RFLP) analysis the primers proposed by Kruse et al (1994) were used to amplify the RNA-3 as described above which was then cut separately with enzymes EcoRI BamHI and MspI 3 Results and discussion31 Detection of BNYVVA total of 9 spinach plants and 25 R raphanistrum samples were collected from the spinach field and tested by DAS-ELISA In this study BNYVV was detected in 20 samples of R raphanistrum collected from the field (Figure 1A) On the other hand 7 of the field-grown spinach samples were infected with BNYVV Generally BNYVV-positive spinach plants have yellow-green mottling symptom on their leaves Although spinach is grown in many regions of Turkey BNYVV has only been recently recorded in spinach production areas in İzmir and Manisa provinces located in the western part of Turkey (Gumus et al 2014) In the ELISA tests with the R raphanistrum samples color development was very slow For this reason the DAS-ELISA test microtiter plates were read following 2 h substrate incubation at room temperature and then again following overnight substrate incubation at 4 degC Even after overnight incubations relatively low A405 values were obtained for most of the samples The positive controls supplied by the Sediag Biochemica and root samples from the sugar beet bait plants grown in BNYVV-infested soils gave satisfactory positive results at 2 h and 16 h substrate incubations The reason for the low ELISA values could be low virus titers in R raphanistrum samples In addition RT-PCR assay to detect BNYVV and P betae was carried out on total RNA extracted from leaves and root samples from R raphanistrum and root samples from the sugar beet bait All samples tested positive for BNYVV

A B

Figure 1 Yellow vein banding in field-grown (A) and greenhouse-grown (B) BNYVV-infected Raphanus raphanistrum

123

KUTLUK YILMAZ et al Turk J Agric For

in this RT-PCR detection assay (Figure 2) Until now BNYVV host studies on Brassicaceae members such as R raphanistrum R sativus B napus and S arvensis have failed to demonstrate infection by this virus or its vector (Barr and Asher 1992 Hugo et al 1996 Legreve et al 2005) However BNYVV and P betae were both detected in Capsella bursa-pastoris (L) Medik and Thlaspi arvense L which belong to the family Brassicaceae in a bait plant test using RT-PCR by Legreve et al (2005) However in this study neither BNYVV nor its vector were found in these species by ELISA or light microscopy respectively (Legreve et al 2005) Later on Mouhanna et al (2008) found that C bursa-pastoris was infested with P betae and BNYVV using bait plant test and back transmission studies To our knowledge this is the first report of a BNYVV infection of R raphanistrum in the family Brassicaceae under natural conditions 32 RFLP analysis of BNYVVKruse et al (1994) showed that RNA-3 of B type strains is cut by BamHI EcoRI StyI and MspI In contrast the A type strains were not cut by any of these restriction enzymes Our BNYVV isolates were not digested by these restriction enzymes suggesting that the isolate from R raphanistrum belongs to the A type strain (Figure 3) The A type the most widespread strain (Schirmer et al 2005) has been previously detected in Turkey (Kruse et al 1994 Kutluk Yilmaz et al 2007) the B type strain is more restricted and mainly found in Germany France (Kruse et al 1994 Koenig et al 2008) Japan (Miyanishi et al 1999) and China (Li et al 2008)33 Detection of P betae in infested rootsRoots of R raphanistrum that were collected from a spinach field were inspected for the presence of P betae using light microscopy P betae cystosori were not detected in roots of any of these plants though some

objects resembling sporogenic plasmodia of P betae were described in the root cells of R raphanistrum (Figure 4) To confirm this result sugar beet bait plant and field-grown R raphanistrum samples were subjected to RT-PCR assay with Polymyxa-specific primers (Ward and Adams 1998) to determine whether they were infected with Polymyxa spp Ward and Adams (1998) emphasized that the products of amplification using these primers on isolates of Polymyxa graminis types I and II and P betae could all be distinguished from one another on 12 agarose gels Therefore a 250-bp DNA fragment expected for the ITS

M 1 2 3 4

750-bp

1000-bp

1500-bp

Figure 2 Confirmation of the presence of BNYVV in leaves of R raphanistrum using RT-PCR analysis M DNA size marker The arrow indicates the BNYVV-specific 1218-bp DNA fragment Lane 1 negative control (water) Lane 2 BNYVV-containing R raphanistrum leaves Lane 3 healthy control (noninfected sugar beet cultivar Arosa) Lane 4 positive control (sugar beet infected with BNYVV)

M 1 2 3 4 5 6 7 8

Figure 3 The results of RFLP analysis of the RT-PCR products of BNYVV RNA-3 M 1-kb ladder (Promega USA) The arrow indicates the BNYVV RNA-3-specific 1218-bp DNA fragment Lanes 1ndash4 RNA from roots of R raphanistrum (EcoR1 BamH1 Msp1 and uncut PCR product) Lanes 5ndash8 RNA from the roots of the bait sugar beet plant grown in BNYVV-infested soil (EcoR1 BamH1 Msp1 and uncut PCR product)

Figure 4 P betae sporogenic plasmodia in root cells of R raphanistrum

124

KUTLUK YILMAZ et al Turk J Agric For

between the 58 s and the 18 s rRNA of P betae was obtained from the roots of the susceptible sugar beet cultivar grown in rhizomania-infested soils and R raphanistrum in this assay (Figure 5) Rysanek et al (2008) suggested that the PCR method would be useful for verifying the presence of P betae in the case of low numbers of cystosori in host range studies Moreover in some plant species plasmodia or zoosporangia could be present without cystosori PCR could reveal such hidden infections Similarly Legreve et al (2008) found that BNYVV and P betae were detected by RT-PCR in some species belonging to different families while the virus and vector went undetected by ELISA and microscopy respectively We suggest that viruliferous P betae zoospores were able to penetrate the roots of R raphanistrum plants and introduce the virus but the protozoa were unable to produce cystosori and complete their life cycle in the roots of wild radish An incomplete infection similar to our result was reported with P betae in Gomphrena globosa (Al Musa and Mink 1981) and P graminis in Arachis hypogaea (groundnut) (Thouvenel and Fauquet 1981) Later on Barr et al (1995) reported similar observations about early P betae developmental stages without cystosori in two wild beet species Beta patellaris and Beta procumbens Although the wild radish leaves became infected naturally with BNYVV in the field they were unable to provide sources of virus infection to other plants whereas comparable sugar beet roots were able to do so There are various factors such as environmental conditions the lack of vital nutritional

factors or insufficient nutrients which might have limited the development of pathogen structures (Barr et al 1995)34 Bait plant and back inoculation testsAmong all Brassicaceae species tested for P betae and BNYVV host status sugar beet spinach Chenopodium album L Chenopodium murale L Amaranthus retroflexus L and Portulaca oleracea L were added to the bait plant test The Table shows the species that became infected with either P betae or BNYVV In this study P betae cystosori were found by light microscopy only in sugar beet C album A retroflexus and P oleracea Interestingly we could not detect either P betae or BNYVV in spinach plants From all tested species infection by BNYVV was only detected in sugar beet and R raphanistrum using ELISA R raphanistrum plants also showed symptoms of yellow vein banding under controlled conditions similar to natural conditions (Figure 1B) However P betae was not detected using the light microscopy technique in these plant roots Neither BNYVV nor P betae was detected in the other tested Brassicaceae species R sativus E sativa B napus B rapa B juncea and B oleracea (Table)

Besides this we tried using roots of a plant infected with a vector or the virus (sugar beet C album A retroflexus P oleracea and R raphanistrum) to see if they had the potential to act as an alternative host for P betae and BNYVV to transmit to susceptible sugar beet plants Transmission back to sugar beet occurred only from B vulgaris (Table)

In rhizomania disease the virusndashvectorndashhost relationships are very complex Over the last decades the existence of different genotypic groups has become obvious and they correspond to differences in pathogenicity and specific geographical region BNYVV isolates fall into three strain groups A B and P types (Kruse et al 1994 Koenig et al 1995 Koenig and Lennefors 2000 Ward et al 2007) In the last years resistance-breaking BNYVV strains have been found in cultivars carrying the Rz1 gene in the United States Europe (Liu and Lewellen 2007 Bornemann et al 2015) and Turkey (Kutluk Yilmaz et al 2013) Beside this the performance of viruliferous P betae in soils is influenced by many different biotic and abiotic factors Gerik and Duffus (1988) described differences in vectoring abilities of P betae populations depending on their origin There are also subgroups of P betae with specific host ranges such as betae amaranthi and the family Portulacaceae (Barr 1979 Abe and Tamada 1986 Abe and Ui 1986) Moreover weeds are populations with a wide range of genetic variability In this respect this appears to be the first report of R raphanistrum from the family Brassicaceae acting as a host of BNYVV under natural conditions The Raphanus isolate was assigned to the A type strain based on RFLP analysis Although P betae cystosori were not found in the roots of

M 1 2 3 4 5

1000- bp

750- bp

500- bp

Figure 5 RT-PCR analysis for detection of P betae M 1-kb DNA ladder (Promega) The arrow indicates the P betae-specific 250-bp DNA fragment Lane 1 root RNA from R raphanistrum Lane 2 negative control (healthy sugar beet roots) Lanes 3ndash5 positive controls (BNYVV-containing sugar beet roots)

125

KUTLUK YILMAZ et al Turk J Agric For

R raphanistrum it may infect wild radish roots without cystosori development Therefore R raphanistrum seems to be a relatively inefficient host for rhizomania compared to sugar beet Nonetheless knowing that this weed is a

host of BNYVV may help to understand its epidemiology and could provide basic information for implementation of effective management strategies Further work is needed in the search for natural hosts of BNYVV and P betae

Table ELISA and microscopy results for the bait plant and back inoculation to B vulgaris tests

Family Plant species Presence of P betae cystosori (microscopy)

Infection byBNYVV (ELISA)

Back inoculation B vulgaris

Chenopodiaceae Beta vulgaris L cv Arosa + + +

Chenopodiaceae Spinacia oleracea L - - nt

Chenopodiaceae Chenopodium album L + - -

Chenopodiaceae Chenopodium murale L - - nt

Amaranthaceae Amaranthus retroflexus L + - -

Portulacaceae Portulaca oleracea L + - -

Brassicaceae Raphanus raphanistrum L - + -

Brassicaceae Raphanus sativus L - - nt

Brassicaceae Brassica napus L (B-12 accession) - - nt

Brassicaceae Brassica napus (B-2 accession) - - nt

Brassicaceae Brassica juncea (L) Czern (B-4 accession) - - nt

Brassicaceae Brassica juncea (B-7 accession) - - nt

Brassicaceae Brassica juncea (B-5 accession) - - nt

Brassicaceae Brassica oleracea L - - nt

Brassicaceae Brassica rapa L - - nt

Brassicaceae Eruca sativa L - - nt

nt not tested

References

Abe H Tamada T (1986) Association of Beet necrotic yellow vein virus with isolates of Polymyxa betae Keskin Ann Phytopathol Soc Jpn 52 235ndash247

Abe H Ui T (1986) Host range of Polymyxa betae Keskin strains in rhizomania-infected soils of sugar beet fields in Japan Ann Phytopathol Soc Jpn 52 394ndash403

Al Musa AM Mink GI (1981) Beet necrotic yellow vein virus in North America Phytopathology 71 773ndash776

Autonel CR Turina M Fantino MG (1995) Spinach virus infections in Italy Inf Fitopatol 45 43ndash47

Barr DJ (1979) Morphology and host range of Polymyxa graminis Polymyxa betae and Ligniera pilorum from Ontario and some other areas Can J Plant Pathol 1 85ndash93

Barr KJ Asher MJC (1992) The host range of Polymyxa betae in Britain Plant Pathol 41 64ndash68

Barr KJ Asher MJC Lewis BG (1995) Resistance to Polymyxa betae in wild Beta species Plant Pathol 44 301ndash307

Blackshaw RE (2001) Influence of wild radish on yield and quality of canola weed Weed Sci 50 344ndash349

Bornemann K Hanse B Varrelmann M Stevens M (2015) Occurrence of resistance-breaking strains of Beet necrotic yellow vein virus in sugar beet in Northwestern Europe and identification of a new variant of the viral pathogenicity factor P25 Plant Pathol 64 25ndash34

Cheam AH Code GR (1995) The biology of Australian weeds 24 Raphanus raphanistrum L Plant Prot Q 10 2ndash13

Clark M Adams AM (1977) Characteristics of microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses J Gen Virol 34 475ndash483

122

KUTLUK YILMAZ et al Turk J Agric For

27 RT-PCR for detection of BNYVV and Polymyxa betaeOne-step RT-PCR was performed as described in the manufacturerrsquos manual (QIAGEN) For the detection of BNYVV the primers proposed by Kruse et al (1994) were used (forward primer GTGATATATATGTGAGGACGCT and reverse primer CCGTGAAATCACGTGTAGTT) which amplified RNA-3 The RT-PCR reaction included the following 155 microL of RNase-free water 1 microL of dNTPs mix (400 microM) 5 microL of 5X Q solution 025 microL each of forward and reverse primers (06 microM) 1 microL of QIAGEN OneStep RT-PCR enzyme mix and 2 microL of RNA sample Reverse transcription consisted of 30 min at 50 degC and 15 min at 95 degC We then ran 35 cycles composed of denaturation for 30 s at 94 degC annealing for 30 s at 53 degC and elongation for 1 min at 72 degC A final elongation of 7 min at 72 degC was also performed

For the detection of Polymyxa in roots of R raphanistrum and sugar beet oligonucleotides flanking 58s-rDNA and internal transcribed spacers (ITS) were used This includes upstream 5`-GAGGCATGCTTCCGAGGGCTCT-3` and downstream primer 5`-CTGCGGAAGGAT-CATTAGCGTT-3`(Ward and Adams 1998) The RT-PCR reaction included the following 152 microL of RNase-free water 1 microL of dNTPs mix (400 microM) 5 microL of 5X Q solution 015 microL each of forward and reverse primers (06 microM) 1 microL of QIAGEN OneStep RT-PCR Enzyme mix and 25 microL of RNA sample Reaction conditions were the same as used for the amplification of RNA-3 except for annealing temperature (55 degC)

After PCR amplification 2 microL of loading buffer was added to the PCR products The samples were analyzed on 12 ethidium bromide agarose gel in 1X Tris-borate-EDTA buffer using the Gel Doc 2000 Systems (Bio-Rad USA) (Ward and Adams 1998)

28 Restriction fragment length polymorphism analysisTo produce a PCR product for restriction fragment length polymorphism (RFLP) analysis the primers proposed by Kruse et al (1994) were used to amplify the RNA-3 as described above which was then cut separately with enzymes EcoRI BamHI and MspI 3 Results and discussion31 Detection of BNYVVA total of 9 spinach plants and 25 R raphanistrum samples were collected from the spinach field and tested by DAS-ELISA In this study BNYVV was detected in 20 samples of R raphanistrum collected from the field (Figure 1A) On the other hand 7 of the field-grown spinach samples were infected with BNYVV Generally BNYVV-positive spinach plants have yellow-green mottling symptom on their leaves Although spinach is grown in many regions of Turkey BNYVV has only been recently recorded in spinach production areas in İzmir and Manisa provinces located in the western part of Turkey (Gumus et al 2014) In the ELISA tests with the R raphanistrum samples color development was very slow For this reason the DAS-ELISA test microtiter plates were read following 2 h substrate incubation at room temperature and then again following overnight substrate incubation at 4 degC Even after overnight incubations relatively low A405 values were obtained for most of the samples The positive controls supplied by the Sediag Biochemica and root samples from the sugar beet bait plants grown in BNYVV-infested soils gave satisfactory positive results at 2 h and 16 h substrate incubations The reason for the low ELISA values could be low virus titers in R raphanistrum samples In addition RT-PCR assay to detect BNYVV and P betae was carried out on total RNA extracted from leaves and root samples from R raphanistrum and root samples from the sugar beet bait All samples tested positive for BNYVV

A B

Figure 1 Yellow vein banding in field-grown (A) and greenhouse-grown (B) BNYVV-infected Raphanus raphanistrum

123

KUTLUK YILMAZ et al Turk J Agric For

in this RT-PCR detection assay (Figure 2) Until now BNYVV host studies on Brassicaceae members such as R raphanistrum R sativus B napus and S arvensis have failed to demonstrate infection by this virus or its vector (Barr and Asher 1992 Hugo et al 1996 Legreve et al 2005) However BNYVV and P betae were both detected in Capsella bursa-pastoris (L) Medik and Thlaspi arvense L which belong to the family Brassicaceae in a bait plant test using RT-PCR by Legreve et al (2005) However in this study neither BNYVV nor its vector were found in these species by ELISA or light microscopy respectively (Legreve et al 2005) Later on Mouhanna et al (2008) found that C bursa-pastoris was infested with P betae and BNYVV using bait plant test and back transmission studies To our knowledge this is the first report of a BNYVV infection of R raphanistrum in the family Brassicaceae under natural conditions 32 RFLP analysis of BNYVVKruse et al (1994) showed that RNA-3 of B type strains is cut by BamHI EcoRI StyI and MspI In contrast the A type strains were not cut by any of these restriction enzymes Our BNYVV isolates were not digested by these restriction enzymes suggesting that the isolate from R raphanistrum belongs to the A type strain (Figure 3) The A type the most widespread strain (Schirmer et al 2005) has been previously detected in Turkey (Kruse et al 1994 Kutluk Yilmaz et al 2007) the B type strain is more restricted and mainly found in Germany France (Kruse et al 1994 Koenig et al 2008) Japan (Miyanishi et al 1999) and China (Li et al 2008)33 Detection of P betae in infested rootsRoots of R raphanistrum that were collected from a spinach field were inspected for the presence of P betae using light microscopy P betae cystosori were not detected in roots of any of these plants though some

objects resembling sporogenic plasmodia of P betae were described in the root cells of R raphanistrum (Figure 4) To confirm this result sugar beet bait plant and field-grown R raphanistrum samples were subjected to RT-PCR assay with Polymyxa-specific primers (Ward and Adams 1998) to determine whether they were infected with Polymyxa spp Ward and Adams (1998) emphasized that the products of amplification using these primers on isolates of Polymyxa graminis types I and II and P betae could all be distinguished from one another on 12 agarose gels Therefore a 250-bp DNA fragment expected for the ITS

M 1 2 3 4

750-bp

1000-bp

1500-bp

Figure 2 Confirmation of the presence of BNYVV in leaves of R raphanistrum using RT-PCR analysis M DNA size marker The arrow indicates the BNYVV-specific 1218-bp DNA fragment Lane 1 negative control (water) Lane 2 BNYVV-containing R raphanistrum leaves Lane 3 healthy control (noninfected sugar beet cultivar Arosa) Lane 4 positive control (sugar beet infected with BNYVV)

M 1 2 3 4 5 6 7 8

Figure 3 The results of RFLP analysis of the RT-PCR products of BNYVV RNA-3 M 1-kb ladder (Promega USA) The arrow indicates the BNYVV RNA-3-specific 1218-bp DNA fragment Lanes 1ndash4 RNA from roots of R raphanistrum (EcoR1 BamH1 Msp1 and uncut PCR product) Lanes 5ndash8 RNA from the roots of the bait sugar beet plant grown in BNYVV-infested soil (EcoR1 BamH1 Msp1 and uncut PCR product)

Figure 4 P betae sporogenic plasmodia in root cells of R raphanistrum

124

KUTLUK YILMAZ et al Turk J Agric For

between the 58 s and the 18 s rRNA of P betae was obtained from the roots of the susceptible sugar beet cultivar grown in rhizomania-infested soils and R raphanistrum in this assay (Figure 5) Rysanek et al (2008) suggested that the PCR method would be useful for verifying the presence of P betae in the case of low numbers of cystosori in host range studies Moreover in some plant species plasmodia or zoosporangia could be present without cystosori PCR could reveal such hidden infections Similarly Legreve et al (2008) found that BNYVV and P betae were detected by RT-PCR in some species belonging to different families while the virus and vector went undetected by ELISA and microscopy respectively We suggest that viruliferous P betae zoospores were able to penetrate the roots of R raphanistrum plants and introduce the virus but the protozoa were unable to produce cystosori and complete their life cycle in the roots of wild radish An incomplete infection similar to our result was reported with P betae in Gomphrena globosa (Al Musa and Mink 1981) and P graminis in Arachis hypogaea (groundnut) (Thouvenel and Fauquet 1981) Later on Barr et al (1995) reported similar observations about early P betae developmental stages without cystosori in two wild beet species Beta patellaris and Beta procumbens Although the wild radish leaves became infected naturally with BNYVV in the field they were unable to provide sources of virus infection to other plants whereas comparable sugar beet roots were able to do so There are various factors such as environmental conditions the lack of vital nutritional

factors or insufficient nutrients which might have limited the development of pathogen structures (Barr et al 1995)34 Bait plant and back inoculation testsAmong all Brassicaceae species tested for P betae and BNYVV host status sugar beet spinach Chenopodium album L Chenopodium murale L Amaranthus retroflexus L and Portulaca oleracea L were added to the bait plant test The Table shows the species that became infected with either P betae or BNYVV In this study P betae cystosori were found by light microscopy only in sugar beet C album A retroflexus and P oleracea Interestingly we could not detect either P betae or BNYVV in spinach plants From all tested species infection by BNYVV was only detected in sugar beet and R raphanistrum using ELISA R raphanistrum plants also showed symptoms of yellow vein banding under controlled conditions similar to natural conditions (Figure 1B) However P betae was not detected using the light microscopy technique in these plant roots Neither BNYVV nor P betae was detected in the other tested Brassicaceae species R sativus E sativa B napus B rapa B juncea and B oleracea (Table)

Besides this we tried using roots of a plant infected with a vector or the virus (sugar beet C album A retroflexus P oleracea and R raphanistrum) to see if they had the potential to act as an alternative host for P betae and BNYVV to transmit to susceptible sugar beet plants Transmission back to sugar beet occurred only from B vulgaris (Table)

In rhizomania disease the virusndashvectorndashhost relationships are very complex Over the last decades the existence of different genotypic groups has become obvious and they correspond to differences in pathogenicity and specific geographical region BNYVV isolates fall into three strain groups A B and P types (Kruse et al 1994 Koenig et al 1995 Koenig and Lennefors 2000 Ward et al 2007) In the last years resistance-breaking BNYVV strains have been found in cultivars carrying the Rz1 gene in the United States Europe (Liu and Lewellen 2007 Bornemann et al 2015) and Turkey (Kutluk Yilmaz et al 2013) Beside this the performance of viruliferous P betae in soils is influenced by many different biotic and abiotic factors Gerik and Duffus (1988) described differences in vectoring abilities of P betae populations depending on their origin There are also subgroups of P betae with specific host ranges such as betae amaranthi and the family Portulacaceae (Barr 1979 Abe and Tamada 1986 Abe and Ui 1986) Moreover weeds are populations with a wide range of genetic variability In this respect this appears to be the first report of R raphanistrum from the family Brassicaceae acting as a host of BNYVV under natural conditions The Raphanus isolate was assigned to the A type strain based on RFLP analysis Although P betae cystosori were not found in the roots of

M 1 2 3 4 5

1000- bp

750- bp

500- bp

Figure 5 RT-PCR analysis for detection of P betae M 1-kb DNA ladder (Promega) The arrow indicates the P betae-specific 250-bp DNA fragment Lane 1 root RNA from R raphanistrum Lane 2 negative control (healthy sugar beet roots) Lanes 3ndash5 positive controls (BNYVV-containing sugar beet roots)

125

KUTLUK YILMAZ et al Turk J Agric For

R raphanistrum it may infect wild radish roots without cystosori development Therefore R raphanistrum seems to be a relatively inefficient host for rhizomania compared to sugar beet Nonetheless knowing that this weed is a

host of BNYVV may help to understand its epidemiology and could provide basic information for implementation of effective management strategies Further work is needed in the search for natural hosts of BNYVV and P betae

Table ELISA and microscopy results for the bait plant and back inoculation to B vulgaris tests

Family Plant species Presence of P betae cystosori (microscopy)

Infection byBNYVV (ELISA)

Back inoculation B vulgaris

Chenopodiaceae Beta vulgaris L cv Arosa + + +

Chenopodiaceae Spinacia oleracea L - - nt

Chenopodiaceae Chenopodium album L + - -

Chenopodiaceae Chenopodium murale L - - nt

Amaranthaceae Amaranthus retroflexus L + - -

Portulacaceae Portulaca oleracea L + - -

Brassicaceae Raphanus raphanistrum L - + -

Brassicaceae Raphanus sativus L - - nt

Brassicaceae Brassica napus L (B-12 accession) - - nt

Brassicaceae Brassica napus (B-2 accession) - - nt

Brassicaceae Brassica juncea (L) Czern (B-4 accession) - - nt

Brassicaceae Brassica juncea (B-7 accession) - - nt

Brassicaceae Brassica juncea (B-5 accession) - - nt

Brassicaceae Brassica oleracea L - - nt

Brassicaceae Brassica rapa L - - nt

Brassicaceae Eruca sativa L - - nt

nt not tested

References

Abe H Tamada T (1986) Association of Beet necrotic yellow vein virus with isolates of Polymyxa betae Keskin Ann Phytopathol Soc Jpn 52 235ndash247

Abe H Ui T (1986) Host range of Polymyxa betae Keskin strains in rhizomania-infected soils of sugar beet fields in Japan Ann Phytopathol Soc Jpn 52 394ndash403

Al Musa AM Mink GI (1981) Beet necrotic yellow vein virus in North America Phytopathology 71 773ndash776

Autonel CR Turina M Fantino MG (1995) Spinach virus infections in Italy Inf Fitopatol 45 43ndash47

Barr DJ (1979) Morphology and host range of Polymyxa graminis Polymyxa betae and Ligniera pilorum from Ontario and some other areas Can J Plant Pathol 1 85ndash93

Barr KJ Asher MJC (1992) The host range of Polymyxa betae in Britain Plant Pathol 41 64ndash68

Barr KJ Asher MJC Lewis BG (1995) Resistance to Polymyxa betae in wild Beta species Plant Pathol 44 301ndash307

Blackshaw RE (2001) Influence of wild radish on yield and quality of canola weed Weed Sci 50 344ndash349

Bornemann K Hanse B Varrelmann M Stevens M (2015) Occurrence of resistance-breaking strains of Beet necrotic yellow vein virus in sugar beet in Northwestern Europe and identification of a new variant of the viral pathogenicity factor P25 Plant Pathol 64 25ndash34

Cheam AH Code GR (1995) The biology of Australian weeds 24 Raphanus raphanistrum L Plant Prot Q 10 2ndash13

Clark M Adams AM (1977) Characteristics of microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses J Gen Virol 34 475ndash483

123

KUTLUK YILMAZ et al Turk J Agric For

in this RT-PCR detection assay (Figure 2) Until now BNYVV host studies on Brassicaceae members such as R raphanistrum R sativus B napus and S arvensis have failed to demonstrate infection by this virus or its vector (Barr and Asher 1992 Hugo et al 1996 Legreve et al 2005) However BNYVV and P betae were both detected in Capsella bursa-pastoris (L) Medik and Thlaspi arvense L which belong to the family Brassicaceae in a bait plant test using RT-PCR by Legreve et al (2005) However in this study neither BNYVV nor its vector were found in these species by ELISA or light microscopy respectively (Legreve et al 2005) Later on Mouhanna et al (2008) found that C bursa-pastoris was infested with P betae and BNYVV using bait plant test and back transmission studies To our knowledge this is the first report of a BNYVV infection of R raphanistrum in the family Brassicaceae under natural conditions 32 RFLP analysis of BNYVVKruse et al (1994) showed that RNA-3 of B type strains is cut by BamHI EcoRI StyI and MspI In contrast the A type strains were not cut by any of these restriction enzymes Our BNYVV isolates were not digested by these restriction enzymes suggesting that the isolate from R raphanistrum belongs to the A type strain (Figure 3) The A type the most widespread strain (Schirmer et al 2005) has been previously detected in Turkey (Kruse et al 1994 Kutluk Yilmaz et al 2007) the B type strain is more restricted and mainly found in Germany France (Kruse et al 1994 Koenig et al 2008) Japan (Miyanishi et al 1999) and China (Li et al 2008)33 Detection of P betae in infested rootsRoots of R raphanistrum that were collected from a spinach field were inspected for the presence of P betae using light microscopy P betae cystosori were not detected in roots of any of these plants though some

objects resembling sporogenic plasmodia of P betae were described in the root cells of R raphanistrum (Figure 4) To confirm this result sugar beet bait plant and field-grown R raphanistrum samples were subjected to RT-PCR assay with Polymyxa-specific primers (Ward and Adams 1998) to determine whether they were infected with Polymyxa spp Ward and Adams (1998) emphasized that the products of amplification using these primers on isolates of Polymyxa graminis types I and II and P betae could all be distinguished from one another on 12 agarose gels Therefore a 250-bp DNA fragment expected for the ITS

M 1 2 3 4

750-bp

1000-bp

1500-bp

Figure 2 Confirmation of the presence of BNYVV in leaves of R raphanistrum using RT-PCR analysis M DNA size marker The arrow indicates the BNYVV-specific 1218-bp DNA fragment Lane 1 negative control (water) Lane 2 BNYVV-containing R raphanistrum leaves Lane 3 healthy control (noninfected sugar beet cultivar Arosa) Lane 4 positive control (sugar beet infected with BNYVV)

M 1 2 3 4 5 6 7 8

Figure 3 The results of RFLP analysis of the RT-PCR products of BNYVV RNA-3 M 1-kb ladder (Promega USA) The arrow indicates the BNYVV RNA-3-specific 1218-bp DNA fragment Lanes 1ndash4 RNA from roots of R raphanistrum (EcoR1 BamH1 Msp1 and uncut PCR product) Lanes 5ndash8 RNA from the roots of the bait sugar beet plant grown in BNYVV-infested soil (EcoR1 BamH1 Msp1 and uncut PCR product)

Figure 4 P betae sporogenic plasmodia in root cells of R raphanistrum

124

KUTLUK YILMAZ et al Turk J Agric For

between the 58 s and the 18 s rRNA of P betae was obtained from the roots of the susceptible sugar beet cultivar grown in rhizomania-infested soils and R raphanistrum in this assay (Figure 5) Rysanek et al (2008) suggested that the PCR method would be useful for verifying the presence of P betae in the case of low numbers of cystosori in host range studies Moreover in some plant species plasmodia or zoosporangia could be present without cystosori PCR could reveal such hidden infections Similarly Legreve et al (2008) found that BNYVV and P betae were detected by RT-PCR in some species belonging to different families while the virus and vector went undetected by ELISA and microscopy respectively We suggest that viruliferous P betae zoospores were able to penetrate the roots of R raphanistrum plants and introduce the virus but the protozoa were unable to produce cystosori and complete their life cycle in the roots of wild radish An incomplete infection similar to our result was reported with P betae in Gomphrena globosa (Al Musa and Mink 1981) and P graminis in Arachis hypogaea (groundnut) (Thouvenel and Fauquet 1981) Later on Barr et al (1995) reported similar observations about early P betae developmental stages without cystosori in two wild beet species Beta patellaris and Beta procumbens Although the wild radish leaves became infected naturally with BNYVV in the field they were unable to provide sources of virus infection to other plants whereas comparable sugar beet roots were able to do so There are various factors such as environmental conditions the lack of vital nutritional

factors or insufficient nutrients which might have limited the development of pathogen structures (Barr et al 1995)34 Bait plant and back inoculation testsAmong all Brassicaceae species tested for P betae and BNYVV host status sugar beet spinach Chenopodium album L Chenopodium murale L Amaranthus retroflexus L and Portulaca oleracea L were added to the bait plant test The Table shows the species that became infected with either P betae or BNYVV In this study P betae cystosori were found by light microscopy only in sugar beet C album A retroflexus and P oleracea Interestingly we could not detect either P betae or BNYVV in spinach plants From all tested species infection by BNYVV was only detected in sugar beet and R raphanistrum using ELISA R raphanistrum plants also showed symptoms of yellow vein banding under controlled conditions similar to natural conditions (Figure 1B) However P betae was not detected using the light microscopy technique in these plant roots Neither BNYVV nor P betae was detected in the other tested Brassicaceae species R sativus E sativa B napus B rapa B juncea and B oleracea (Table)

Besides this we tried using roots of a plant infected with a vector or the virus (sugar beet C album A retroflexus P oleracea and R raphanistrum) to see if they had the potential to act as an alternative host for P betae and BNYVV to transmit to susceptible sugar beet plants Transmission back to sugar beet occurred only from B vulgaris (Table)

In rhizomania disease the virusndashvectorndashhost relationships are very complex Over the last decades the existence of different genotypic groups has become obvious and they correspond to differences in pathogenicity and specific geographical region BNYVV isolates fall into three strain groups A B and P types (Kruse et al 1994 Koenig et al 1995 Koenig and Lennefors 2000 Ward et al 2007) In the last years resistance-breaking BNYVV strains have been found in cultivars carrying the Rz1 gene in the United States Europe (Liu and Lewellen 2007 Bornemann et al 2015) and Turkey (Kutluk Yilmaz et al 2013) Beside this the performance of viruliferous P betae in soils is influenced by many different biotic and abiotic factors Gerik and Duffus (1988) described differences in vectoring abilities of P betae populations depending on their origin There are also subgroups of P betae with specific host ranges such as betae amaranthi and the family Portulacaceae (Barr 1979 Abe and Tamada 1986 Abe and Ui 1986) Moreover weeds are populations with a wide range of genetic variability In this respect this appears to be the first report of R raphanistrum from the family Brassicaceae acting as a host of BNYVV under natural conditions The Raphanus isolate was assigned to the A type strain based on RFLP analysis Although P betae cystosori were not found in the roots of

M 1 2 3 4 5

1000- bp

750- bp

500- bp

Figure 5 RT-PCR analysis for detection of P betae M 1-kb DNA ladder (Promega) The arrow indicates the P betae-specific 250-bp DNA fragment Lane 1 root RNA from R raphanistrum Lane 2 negative control (healthy sugar beet roots) Lanes 3ndash5 positive controls (BNYVV-containing sugar beet roots)

125

KUTLUK YILMAZ et al Turk J Agric For

R raphanistrum it may infect wild radish roots without cystosori development Therefore R raphanistrum seems to be a relatively inefficient host for rhizomania compared to sugar beet Nonetheless knowing that this weed is a

host of BNYVV may help to understand its epidemiology and could provide basic information for implementation of effective management strategies Further work is needed in the search for natural hosts of BNYVV and P betae

Table ELISA and microscopy results for the bait plant and back inoculation to B vulgaris tests

Family Plant species Presence of P betae cystosori (microscopy)

Infection byBNYVV (ELISA)

Back inoculation B vulgaris

Chenopodiaceae Beta vulgaris L cv Arosa + + +

Chenopodiaceae Spinacia oleracea L - - nt

Chenopodiaceae Chenopodium album L + - -

Chenopodiaceae Chenopodium murale L - - nt

Amaranthaceae Amaranthus retroflexus L + - -

Portulacaceae Portulaca oleracea L + - -

Brassicaceae Raphanus raphanistrum L - + -

Brassicaceae Raphanus sativus L - - nt

Brassicaceae Brassica napus L (B-12 accession) - - nt

Brassicaceae Brassica napus (B-2 accession) - - nt

Brassicaceae Brassica juncea (L) Czern (B-4 accession) - - nt

Brassicaceae Brassica juncea (B-7 accession) - - nt

Brassicaceae Brassica juncea (B-5 accession) - - nt

Brassicaceae Brassica oleracea L - - nt

Brassicaceae Brassica rapa L - - nt

Brassicaceae Eruca sativa L - - nt

nt not tested

References

Abe H Tamada T (1986) Association of Beet necrotic yellow vein virus with isolates of Polymyxa betae Keskin Ann Phytopathol Soc Jpn 52 235ndash247

Abe H Ui T (1986) Host range of Polymyxa betae Keskin strains in rhizomania-infected soils of sugar beet fields in Japan Ann Phytopathol Soc Jpn 52 394ndash403

Al Musa AM Mink GI (1981) Beet necrotic yellow vein virus in North America Phytopathology 71 773ndash776

Autonel CR Turina M Fantino MG (1995) Spinach virus infections in Italy Inf Fitopatol 45 43ndash47

Barr DJ (1979) Morphology and host range of Polymyxa graminis Polymyxa betae and Ligniera pilorum from Ontario and some other areas Can J Plant Pathol 1 85ndash93

Barr KJ Asher MJC (1992) The host range of Polymyxa betae in Britain Plant Pathol 41 64ndash68

Barr KJ Asher MJC Lewis BG (1995) Resistance to Polymyxa betae in wild Beta species Plant Pathol 44 301ndash307

Blackshaw RE (2001) Influence of wild radish on yield and quality of canola weed Weed Sci 50 344ndash349

Bornemann K Hanse B Varrelmann M Stevens M (2015) Occurrence of resistance-breaking strains of Beet necrotic yellow vein virus in sugar beet in Northwestern Europe and identification of a new variant of the viral pathogenicity factor P25 Plant Pathol 64 25ndash34

Cheam AH Code GR (1995) The biology of Australian weeds 24 Raphanus raphanistrum L Plant Prot Q 10 2ndash13

Clark M Adams AM (1977) Characteristics of microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses J Gen Virol 34 475ndash483

124

KUTLUK YILMAZ et al Turk J Agric For

between the 58 s and the 18 s rRNA of P betae was obtained from the roots of the susceptible sugar beet cultivar grown in rhizomania-infested soils and R raphanistrum in this assay (Figure 5) Rysanek et al (2008) suggested that the PCR method would be useful for verifying the presence of P betae in the case of low numbers of cystosori in host range studies Moreover in some plant species plasmodia or zoosporangia could be present without cystosori PCR could reveal such hidden infections Similarly Legreve et al (2008) found that BNYVV and P betae were detected by RT-PCR in some species belonging to different families while the virus and vector went undetected by ELISA and microscopy respectively We suggest that viruliferous P betae zoospores were able to penetrate the roots of R raphanistrum plants and introduce the virus but the protozoa were unable to produce cystosori and complete their life cycle in the roots of wild radish An incomplete infection similar to our result was reported with P betae in Gomphrena globosa (Al Musa and Mink 1981) and P graminis in Arachis hypogaea (groundnut) (Thouvenel and Fauquet 1981) Later on Barr et al (1995) reported similar observations about early P betae developmental stages without cystosori in two wild beet species Beta patellaris and Beta procumbens Although the wild radish leaves became infected naturally with BNYVV in the field they were unable to provide sources of virus infection to other plants whereas comparable sugar beet roots were able to do so There are various factors such as environmental conditions the lack of vital nutritional

factors or insufficient nutrients which might have limited the development of pathogen structures (Barr et al 1995)34 Bait plant and back inoculation testsAmong all Brassicaceae species tested for P betae and BNYVV host status sugar beet spinach Chenopodium album L Chenopodium murale L Amaranthus retroflexus L and Portulaca oleracea L were added to the bait plant test The Table shows the species that became infected with either P betae or BNYVV In this study P betae cystosori were found by light microscopy only in sugar beet C album A retroflexus and P oleracea Interestingly we could not detect either P betae or BNYVV in spinach plants From all tested species infection by BNYVV was only detected in sugar beet and R raphanistrum using ELISA R raphanistrum plants also showed symptoms of yellow vein banding under controlled conditions similar to natural conditions (Figure 1B) However P betae was not detected using the light microscopy technique in these plant roots Neither BNYVV nor P betae was detected in the other tested Brassicaceae species R sativus E sativa B napus B rapa B juncea and B oleracea (Table)

Besides this we tried using roots of a plant infected with a vector or the virus (sugar beet C album A retroflexus P oleracea and R raphanistrum) to see if they had the potential to act as an alternative host for P betae and BNYVV to transmit to susceptible sugar beet plants Transmission back to sugar beet occurred only from B vulgaris (Table)

In rhizomania disease the virusndashvectorndashhost relationships are very complex Over the last decades the existence of different genotypic groups has become obvious and they correspond to differences in pathogenicity and specific geographical region BNYVV isolates fall into three strain groups A B and P types (Kruse et al 1994 Koenig et al 1995 Koenig and Lennefors 2000 Ward et al 2007) In the last years resistance-breaking BNYVV strains have been found in cultivars carrying the Rz1 gene in the United States Europe (Liu and Lewellen 2007 Bornemann et al 2015) and Turkey (Kutluk Yilmaz et al 2013) Beside this the performance of viruliferous P betae in soils is influenced by many different biotic and abiotic factors Gerik and Duffus (1988) described differences in vectoring abilities of P betae populations depending on their origin There are also subgroups of P betae with specific host ranges such as betae amaranthi and the family Portulacaceae (Barr 1979 Abe and Tamada 1986 Abe and Ui 1986) Moreover weeds are populations with a wide range of genetic variability In this respect this appears to be the first report of R raphanistrum from the family Brassicaceae acting as a host of BNYVV under natural conditions The Raphanus isolate was assigned to the A type strain based on RFLP analysis Although P betae cystosori were not found in the roots of

M 1 2 3 4 5

1000- bp

750- bp

500- bp

Figure 5 RT-PCR analysis for detection of P betae M 1-kb DNA ladder (Promega) The arrow indicates the P betae-specific 250-bp DNA fragment Lane 1 root RNA from R raphanistrum Lane 2 negative control (healthy sugar beet roots) Lanes 3ndash5 positive controls (BNYVV-containing sugar beet roots)

125

KUTLUK YILMAZ et al Turk J Agric For

R raphanistrum it may infect wild radish roots without cystosori development Therefore R raphanistrum seems to be a relatively inefficient host for rhizomania compared to sugar beet Nonetheless knowing that this weed is a

host of BNYVV may help to understand its epidemiology and could provide basic information for implementation of effective management strategies Further work is needed in the search for natural hosts of BNYVV and P betae

Table ELISA and microscopy results for the bait plant and back inoculation to B vulgaris tests

Family Plant species Presence of P betae cystosori (microscopy)

Infection byBNYVV (ELISA)

Back inoculation B vulgaris

Chenopodiaceae Beta vulgaris L cv Arosa + + +

Chenopodiaceae Spinacia oleracea L - - nt

Chenopodiaceae Chenopodium album L + - -

Chenopodiaceae Chenopodium murale L - - nt

Amaranthaceae Amaranthus retroflexus L + - -

Portulacaceae Portulaca oleracea L + - -

Brassicaceae Raphanus raphanistrum L - + -

Brassicaceae Raphanus sativus L - - nt

Brassicaceae Brassica napus L (B-12 accession) - - nt

Brassicaceae Brassica napus (B-2 accession) - - nt

Brassicaceae Brassica juncea (L) Czern (B-4 accession) - - nt

Brassicaceae Brassica juncea (B-7 accession) - - nt

Brassicaceae Brassica juncea (B-5 accession) - - nt

Brassicaceae Brassica oleracea L - - nt

Brassicaceae Brassica rapa L - - nt

Brassicaceae Eruca sativa L - - nt

nt not tested

References

Abe H Tamada T (1986) Association of Beet necrotic yellow vein virus with isolates of Polymyxa betae Keskin Ann Phytopathol Soc Jpn 52 235ndash247

Abe H Ui T (1986) Host range of Polymyxa betae Keskin strains in rhizomania-infected soils of sugar beet fields in Japan Ann Phytopathol Soc Jpn 52 394ndash403

Al Musa AM Mink GI (1981) Beet necrotic yellow vein virus in North America Phytopathology 71 773ndash776

Autonel CR Turina M Fantino MG (1995) Spinach virus infections in Italy Inf Fitopatol 45 43ndash47

Barr DJ (1979) Morphology and host range of Polymyxa graminis Polymyxa betae and Ligniera pilorum from Ontario and some other areas Can J Plant Pathol 1 85ndash93

Barr KJ Asher MJC (1992) The host range of Polymyxa betae in Britain Plant Pathol 41 64ndash68

Barr KJ Asher MJC Lewis BG (1995) Resistance to Polymyxa betae in wild Beta species Plant Pathol 44 301ndash307

Blackshaw RE (2001) Influence of wild radish on yield and quality of canola weed Weed Sci 50 344ndash349

Bornemann K Hanse B Varrelmann M Stevens M (2015) Occurrence of resistance-breaking strains of Beet necrotic yellow vein virus in sugar beet in Northwestern Europe and identification of a new variant of the viral pathogenicity factor P25 Plant Pathol 64 25ndash34

Cheam AH Code GR (1995) The biology of Australian weeds 24 Raphanus raphanistrum L Plant Prot Q 10 2ndash13

Clark M Adams AM (1977) Characteristics of microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses J Gen Virol 34 475ndash483

125

KUTLUK YILMAZ et al Turk J Agric For

R raphanistrum it may infect wild radish roots without cystosori development Therefore R raphanistrum seems to be a relatively inefficient host for rhizomania compared to sugar beet Nonetheless knowing that this weed is a

host of BNYVV may help to understand its epidemiology and could provide basic information for implementation of effective management strategies Further work is needed in the search for natural hosts of BNYVV and P betae

Table ELISA and microscopy results for the bait plant and back inoculation to B vulgaris tests

Family Plant species Presence of P betae cystosori (microscopy)

Infection byBNYVV (ELISA)

Back inoculation B vulgaris

Chenopodiaceae Beta vulgaris L cv Arosa + + +

Chenopodiaceae Spinacia oleracea L - - nt

Chenopodiaceae Chenopodium album L + - -

Chenopodiaceae Chenopodium murale L - - nt

Amaranthaceae Amaranthus retroflexus L + - -

Portulacaceae Portulaca oleracea L + - -

Brassicaceae Raphanus raphanistrum L - + -

Brassicaceae Raphanus sativus L - - nt

Brassicaceae Brassica napus L (B-12 accession) - - nt

Brassicaceae Brassica napus (B-2 accession) - - nt

Brassicaceae Brassica juncea (L) Czern (B-4 accession) - - nt

Brassicaceae Brassica juncea (B-7 accession) - - nt

Brassicaceae Brassica juncea (B-5 accession) - - nt

Brassicaceae Brassica oleracea L - - nt

Brassicaceae Brassica rapa L - - nt

Brassicaceae Eruca sativa L - - nt

nt not tested

References

Abe H Tamada T (1986) Association of Beet necrotic yellow vein virus with isolates of Polymyxa betae Keskin Ann Phytopathol Soc Jpn 52 235ndash247

Abe H Ui T (1986) Host range of Polymyxa betae Keskin strains in rhizomania-infected soils of sugar beet fields in Japan Ann Phytopathol Soc Jpn 52 394ndash403

Al Musa AM Mink GI (1981) Beet necrotic yellow vein virus in North America Phytopathology 71 773ndash776

Autonel CR Turina M Fantino MG (1995) Spinach virus infections in Italy Inf Fitopatol 45 43ndash47

Barr DJ (1979) Morphology and host range of Polymyxa graminis Polymyxa betae and Ligniera pilorum from Ontario and some other areas Can J Plant Pathol 1 85ndash93

Barr KJ Asher MJC (1992) The host range of Polymyxa betae in Britain Plant Pathol 41 64ndash68

Barr KJ Asher MJC Lewis BG (1995) Resistance to Polymyxa betae in wild Beta species Plant Pathol 44 301ndash307

Blackshaw RE (2001) Influence of wild radish on yield and quality of canola weed Weed Sci 50 344ndash349

Bornemann K Hanse B Varrelmann M Stevens M (2015) Occurrence of resistance-breaking strains of Beet necrotic yellow vein virus in sugar beet in Northwestern Europe and identification of a new variant of the viral pathogenicity factor P25 Plant Pathol 64 25ndash34

Cheam AH Code GR (1995) The biology of Australian weeds 24 Raphanus raphanistrum L Plant Prot Q 10 2ndash13

Clark M Adams AM (1977) Characteristics of microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses J Gen Virol 34 475ndash483

126

KUTLUK YILMAZ et al Turk J Agric For

Erdiller G Oumlzguumlr OE (1994) Rhizomania diseases of sugar beet in Turkey In Proceedings of the 9th Congress of the Mediterranean Phytopathological Union 18ndash24 September 1994 Kuşadası Turkey Florence Italy Mediterranean Phytopathological Union pp 443ndash446

Gerik JS Duffus JE (1988) Differences in vectoring ability and aggressiveness of isolates of Polymyxa betae Phytopathology 78 1340ndash1343

Gumus M Erbay E Erkan S Paylan IC (2014) Occurrence of viruses infecting spinach in Western Anatolia of Turkey the first field survey report J Food Agric Environ 12 272ndash275

Hugo SA Henry CM Harju V (1996) The role of alternative hosts of Polymyxa betae in transmission of Beet necrotic yellow vein virus (BNYVV) in England J Plant Pathol 45 662ndash666

Kaya R (2009) Distribution of rhizomania disease in sugar beet growing areas of Turkey J Agric Sci 15 332ndash340

Koch F (1987) Bericht uumlber eine in verschiedene zuckerruumlbenanbaugebiete der Turkseker in Anatolien und Thrazien zum stadium von wurzelerkrankungen Einbeck Germany KWS Kleinwanzlebener Saatzucht (in German)

Koenig R Kastirr U Holtschulte B Deml G Varrelmann M (2008) Distribution of various types and P25 subtypes of Beet necrotic yellow vein virus in Germany and other European countries Arch Virol 153 2139ndash2144

Koenig R Lennefors BL (2000) Molecular analyses of European A B and P type sources of Beet necrotic yellow vein virus and detection of the rare P type in Kazakhstan Arch Virol 145 1561ndash1570

Koenig R Luumlddecke P Haeberle AM (1995) Detection of beet necrotic yellow vein virus strains variants and mixed infections by examining single-strand conformation polymorphisms of immunocapture RT-PCR products J Gen Virol 76 2051ndash2055

Koenig R Pleij CWA Beier C Commandeur U (1998) Genome properties of beet virus Q a new furo-like virus from sugar beet determined from unpurified virus J Gen Virol 79 2027ndash2036

Kruse M Koenig R Hoffman A Kaufmann A Commandeur U Solevyev AG Savenkov I Burgermeister W (1994) Restriction fragment length polymorphism analysis of reverse transcription-PCR products reveals the existence of two major strain groups of beet necrotic yellow vein virus J Gen Virol 75 1835ndash1842

Kutluk Yilmaz ND Arli Sokmen M (2010) Occurrences of sugar beet soilborne viruses transmitted by Polymyxa betae northern and central Turkey J Plant Pathol 92 507ndash510

Kutluk Yilmaz ND Arli Sokmen M Kaya R Toksoz Y Tunali B Erkan E Sevik MA (2012) Resistance-breaking isolates of Beet necrotic yellow vein virus (BNYVV) collected from sugar beet fields in the northern eastern and central parts of Turkey In Proceedings of the 10th Conference of the European Foundation for Plant Pathology ndash IPM 20 Towards Future-Proof Crop Protection in Europe 1ndash5 October 2012 Wageningen the Netherlands Wageningen the Netherlands EFPP p 39

Kutluk Yilmaz ND Meunier A Schmit JF Stas A Bragard C (2007) Partial nucleotide sequence analysis of Turkish isolates of Beet necrotic yellow vein virus (BNYVV) RNA-3 J Plant Pathol 56 311ndash316

Legreve A Schmit JF Bragard C Maraite H (2005) The role of climate and alternative hosts in the epidemiology of rhizomania In Rush CM editor Proceedings of the Sixth Symposium of the International Working Group on Plant Viruses with Fungal Vectors 5ndash7 September 2005 Bologna Italy Zurich Switzerland IWGPVFV pp 125ndash128

Li M Liu T Wang B Han CG Li DW Yu JL (2008) Phylogenetic analysis of Beet necrotic yellow vein virus isolates from China Virus Genes 36 429ndash432

Liu HY Lewellen RT (2007) Distribution and molecular characterization of resistance-breaking isolates of Beet necrotic yellow vein virus in the United States Plant Dis 91 847ndash851

Liu HY Mou B Richardson K Koike ST (2010) First report of Beet necrotic yellow vein virus infecting spinach in California Plant Dis 94 640

Meunier A Schmit JF Stas A Kutluk N Bragard C (2003) Multiplex reverse transcription for simultaneous detection of Beet necrotic yellow vein virus Beet soilborne virus and Beet virus Q and their vector Polymyxa betae Keskin on sugar beet Appl Environ Microbiol 69 2356ndash2360

Miyanishi M Kusume T Saito M Tamada T (1999) Evidence for three groups of sequence variants of beet necrotic yellow vein virus RNA 5 Arch Virol 144 879ndash892

Mouhanna AM Langen G Schloumlsser E (2008) Weeds as alternative hosts for BSBV BNYVV and the vector Polymyxa betae (German isolate) J Plant Dis Protect 115 193ndash198

Rysanek P Homa I Zouhar M Rozo M Putz C (2008) Host range and host (un)specificity of different Polymyxa betae isolates In Rush CM editor Proceedings of the Seventh Symposium of the International Working Group on Plant Viruses with Fungal Vectors 1ndash4 September 2008 Quedlinburg Germany Zurich Switzerland IWGPVFV pp 54ndash58

Schirmer A Link D Cognat V Beuve M Meunier A Bragard C Gilmer D Lemaire O (2005) Phylogenetic analysis of isolates of Beet necrotic yellow vein virus collected worldwide J Gen Virol 86 2897ndash2911

Tamada T Baba T (1973) Beet Necrotic Yellow Vein Virus from ldquorhizomaniardquo affected sugar beet in Japan Ann Phytopathol Soc Jpn 39 325ndash332

Thouvenel JC Fauquet C (1981) Further properties of peanut clump virus and studies on its natural transmission An Appl Biol 97 99ndash107

Ward E Adams MJC (1998) Analysis of ribosomal DNA sequences of Polymyxa species and related fungi and the development of genus- and species specific PCR primers Mycol Res 102 965ndash974

Ward L Koenig R Budge G Garrido C McGrath C Stubbey H Boonham N (2007) Occurrence of two different types of RNA-5 containing beet necrotic yellow vein virus in the UK Arc Virol 152 59ndash73