Original research articles

Development of rapid direct PCR assays to identify downy and powdery mildew and grey mould in Vitis vinifera tissues


Aims: The development of a rapid and reliable direct PCR method to detect fungal propagules in grapevine tissues without prior DNA purification steps, and illustration of its potential use with different examples.

Methods and results: Different grapevine samples crushed in the presence of polyvinylpolypyrrolidone (PVPP) were used as templates for direct PCR amplification with primers specifie for Erysiphe necator, Plasmopara viticola, Botrytis cinerea and Vitis vinifera. Sequencing of the PCR products confirmed the specificity of the amplifications. The sensitivity tested using conidia/sporangia dilution series was high, ranging from five sporangia for P. viticola to one conidium for E. necator. The potential of this technique is illustrated through the study of four epidemiological questions. Fungal propagules were observed in dormant buds using microscopy, but the responsible species could not be identified. Direct PCR revealed the presence of E. necator and B. cinerea in 29 % and 65 % of the buds, respectively. Downy mildew could be detected in asymptomatic leaves sampled in fields after potentially infectious events. In bunch, microscopic analysis of rachis sections showed the presence of hyphae growing in the green tissue. Direct PCR identified the presence of P. viticola.

Conclusion: A direct PCR method without DNA purification was demonstrated to be a simple and reliable method for the detection and identification of fungal pathogens in grapevine tissues. This method, together with microscopy, is a very interesting tool that can be used to study various epidemiological problems in the grapevine, including important unanswered questions such as the route of infection that leads to brown rot caused by downy mildew.

Significance and impact of the study: Direct PCR was shown to be a simple and versatile technique for the study of epidemiological questions in the grapevine. This technique could be extended to other pathosystems with minor adaptations.


The study of complex relationships between organisms, such as microorganisms, and their substrates, environment or one another can currently be accomplished using methods that are completely independent of classical culture techniques. Methods such as polymerase chain reaction (PCR) (van Belkum et al., 1998; Sachse, 2004), restriction fragment length polymorphism (RFLP) (Vaneechoutte, 1996), pyrosequencing (Petrosino et al., 2009) and, more recently, metagenomics (Miller et al., 2013; Ross-Davis et al., 2013), metatranscriptomics (McGrath et al., 2010) and other meta’omics methods (Lepage et al., 2013) represent new ways to study complex ecosystems and their interactions (Segata et al., 2013). The development of such methods has enabled the detection of specific organisms within a complex matrix.

PCR has become a widely used technique with applications in all biological and medical fields. However, in most cases, PCR involves prior purification of nucleic acids. A direct PCR method, without any DNA purification steps, was previously developed to detect latent Botrytis cinerea in young grapevine berries (Gindro et al., 2005). This method, combined with microscopic analyses (Keller et al., 2003), enabled the study of the infection at the bloom stage and during the following latency period. These data provided new insights into the epidemiology of grey mould and confirmed the importance of protecting the grapevine from B. cinerea infections during the bloom stage.

Among the approximately ten species of fungi that are considered to be major grapevine pathogens, three are responsible for most damage in vineyards: grey mould (B. cinerea), downy mildew (Plasmopara viticola) and powdery mildew (Erysiphe necator). A specific problem lies in the obligate biotrophic nature of P. viticola and E. necator, which prohibits their cultivation on artificial media, making their study much more complicated and time consuming. PCR amplification methods have been described for the identification or quantification of B. cinerea in air samples (Carisse et al., 2009), E. necator in wine grapes and must (Stummer et al., 2006) and air samples (Thiessen et al., 2013), and P. viticola in grapevine leaves (Valsesia et al., 2005). Several studies have successfully developed PCR reactions without prior DNA purification, relying upon lysis at the high temperature of amplification. For example, these techniques have been applied to whole bacterial cells (Gussow and Clackson, 1989), tobacco leaves and root pieces (Berthomieu and Meyer, 1991), fungal spores (Aufauvre-Brown et al., 1993) and in direct diagnostics of bacteria and viruses in animal tissues (Olive, 1989). However, it is known that the amplification reaction can be inhibited by different parameters such as excessively high cell numbers, cell wall fragments, DNA-binding proteins, polysaccharides, phenols, detergents or insufficient cell lysis (Moreira, 1998; Dewey and Yohalem, 2004). Plant tissues are very rich in polysaccharides and polyphenols, including secondary metabolites as phytoalexins and lignin (Taiz and Zeiger, 2010), which can potentially inhibit PCR reactions, making purification steps of nucleic acids usually indispensable.

This paper describes a direct PCR method to detect fungal DNA in plants without prior DNA purification steps. The potential of this technique is illustrated through the study of four epidemiological questions concerning grapevines: the presence of E. necator in dormant buds, the occurrence of B. cinerea in dormant buds, the early detection of P. viticola in asymptomatic leaves in the field, and the detection of systemic development of P. viticola in the bunch.

Materials and methods

1. Plant material and artificial infections

Vitis vinifera L. cv. Chasselas grafted on 3309 were obtained from the Agroscope in Changins (Switzerland) and planted in experimental fields in 2005. They were cultivated for eight years until sampling. The experimental plot consisted of one row (distance between rows: 1.5 m; between plants within a row: 0.75 m) conducted in “Guyot”. Leaves, inflorescences, bunches and buds were all sampled from this experimental plot. For P. viticola, the leaves and grape clusters of three plants were artificially infected with an aqueous sporangia suspension at stage BBCH 57 (Hack et al., 1992), namely when the inflorescences are fully developed, as previously described (Gindro et al., 2003). In the case of B. cinerea, artificial infections were realised at the end of bloom (BBCH 69, all caps fallen) with an aqueous conidia suspension as described before (Viret et al., 2004). In the case of E. necator, conidia were collected from infected leaves as described previously (Schnee et al., 2008) and dormant buds were collected in February 2013 from plants in an untreated plot that were strongly infected in 2012. For the early detection of P. viticola infections on V. vinifera cv. Cabernet franc, entire leaves were randomly sampled in a vineyard in Bordeaux the day after potentially infectious conditions (rain or dew) and directly stored at -80°C until use.

2. Sample preparation and PCR amplification

Plant samples, leaves (3 mg), flowers (40 mg), rachis sections (40 mg) and buds (20 mg) were crushed with a plastic pestle in an Eppendorf in 100 ml of water containing 1% (w/v) polyvinylpolypyrrolidone (PVPP), except for buds, which received 500 ml of water. This crude extract was diluted 10- and 100-fold with nanopure water and used directly for PCR. Alternatively, bigger samples, such as entire leaves, were placed into an ELISA bag (Bioreba AG), frozen in liquid nitrogen and crushed with an homogeniser (Homex 6, Bioreba AG) into a fine powder and put on ice or stored at -20°C until use. The powder was then suspended in 2 ml of 1x PBS containing Tween 20 (0.05% v/v) and PVPP (2% w/v). Aliquots of this solution were diluted 10- and 100-fold into water and used directly for PCR amplification. PCR reactions were performed in 25 ml volumes containing 2 units of Taq (Qiagen Taq DNA polymerase), 1x PCR buffer, 0.4 mM of each primer, 0.2 mM of each dNTP, 3 mM MgCl2 and 18.3 ml of the diluted crude extract. The specific primers used in this analysis are described in Table 1. Amplifications were performed in a Biometra T3000 thermocycler with a first step at 97°C for 15 min to lyse the cells, followed by 36 cycles at 94°C for 30 s, 54°C for 30 s, 72°C for 90 s and a final extension step at 72°C for 10 min. Gel electrophoresis of PCR products was performed on 1% agarose gels. Control amplifications were performed on fungal spores or sporangia as described previously (Gindro et al., 2005). Spores and sporangia dilution series (1’000, 100, 10, 1 sporangia or conidia ml-1) were performed to determine the sensitivity of the PCR reaction. Spore suspensions were counted under the microscope on standardised examination chambers (KOVA Glasstic Slide 10 with grids, Hycor Agilent Technologies, Amstelveen, The Netherlands) and the concentration adjusted to 1’000 spores ml-1. PCR products were purified and sequenced to confirm the specificity of the reaction.

Table 1. Sequences of the different primers used in this study and amplicon length.

Organism Primer name Sequence Amplicon (bp)
Plasmopara viticola LSU 1+_PV TAGTAACGGCGAGTGAAGCG 698
Botrytis cinerea C729+ AGCTCGAGAGAGATCTCTGA 730
Vitis vinifera Vvin-F CCTTCAGGTGGGTACAGTGG 241

3. Transmission electron microscopy

Each sample, leaf fragment, rachis, dormant bud fragment and flower fragment was prepared according to Roland and Vian (1991), pre-fixed with a solution of 3% glutaraldehyde-2% paraformaldehyde in 0.07 M phosphate buffer at pH 7, embedded in 2% agarose and post-fixed with a solution of 1% OsO4. The samples were then dehydrated in a graded series of ethanol solutions of 30-50-70-95-100% (v/v) and embedded in LR White resin (14381-UC London Resin Company). After polymerisation (24 h at 60°C), semi-thin (0.8 mm) and thin (0.08 µm) sections were cut and stained with a solution of 1% methylene blue, sodium tetraborate and azure II for the semi-thin sections, or 2% uranyl acetate followed by lead citrate according to Reynolds (1963) for the thin sections. Semi-thin sections were observed using a light microscope (Leica DMLB) equipped with a Leica DFC 490 FX camera. Thin sections were observed with a transmission electron microscope (Philips CM10) with a Mega View II camera.

Results and discussion

Direct PCR is a simple and rapid method for the detection and identification of a pathogen in non-symptomatic plant tissues. Figure 1 shows that the direct PCR methods described here are able to specifically detect the three major fungal pathogens of grapevine from a mix of symptomless infected green tissues. The different bands were purified from the gel and sequenced to confirm the specificity of the PCR amplification. Grapevine green tissues contain many phenolic and polyphenolic compounds as well as sugars, which are all potential inhibitors of PCR reactions. In the case of classical plant DNA extraction, all of these constituents are carefully removed by different purification steps. However, in direct PCR reactions, no prior purification steps are performed, and the inhibitors may still be present in the reaction mix. Some studies summarised the various compounds found in samples that inhibit or facilitate the enzymatic amplification reactions (Rossen et al., 1992; Wilson, 1997). In the case of direct PCR on grapevine green tissue, PVPP, which chelates polyphenolic compounds, is added to the mix prior to the cell lysis step to avoid inhibition (95°C 15 min). For the same purpose, the initial template mixture was diluted 10- to 100-fold. Because of this very rough purification step, it is very important to have a positive control to exclude the possibility that the absence of a band is due to the presence of inhibitors. Each sample was also PCR-amplified with V. vinifera primers as a positive control. The sensitivity of the direct PCR was tested with dilution series of conidia and sporangia in water. The results show a detection threshold of one conidium per microliter for the B. cinerea primers, which is consistent with previous results (Gindro et al., 2005), one sporangium per microliter for P. viticola, and one conidium per microliter for E. necator (Figure 2).

Figure 1. Direct PCR amplification of a mix of grapevine leaf pieces infected with Plasmopara viticola and Erysiphe necator and flowers infected with Botrytis cinerea (infected tissues). Lanes 1 and 14: 1 kb DNA Ladder. Lane 2: P. viticola primers with infected tissues, lane 3: P. viticola primers with healthy tissues, lane 5: E. necator primers with infected tissues, lane 6: E. necator primers with healthy tissues, lane 8: B. cinerea primers with infected tissues, lane 9: B. cinerea primers with healthy tissues, lane 11: V. vinifera primers with infected tissues, lane 12: V. vinifera primers with healthy tissues, lanes 4, 7, 10 and 13: negative control with water. Amplicon sizes are: P. viticola 698 bp, E. necator 379 bp, B. cinerea 730 bp and V. vinifera 241 bp.

Figure 2. Direct PCR sensitivity tests by amplification of Plasmopara viticola sporangia, Erysiphe necator and Botrytis cinerea conidia dilution series. Lanes 1, 8 and 15: 1 kb DNA Ladder. For P. viticola: Lanes 2 to 5: dilution series from 1’000, 100, 10 to 1 sporangia per microliter. Lane 6: negative control with water. Lane 7: empty. For E. necator: Lanes 9 to 12: dilution series from 1’000, 100, 10 to 1 conidia per microliter. Lane 13: negative control with water. Lane 14: empty. For B. cinerea: Lanes 16 to 19: dilution series from 1’000, 100, 10 to 1 conidia per microliter. Lane 20: negative control with water.

The direct PCR method can be used to study different epidemiological questions such as the latency of these pathogens in apparently healthy tissues, or overwintering of pathogens, especially obligate biotrophs, in dormant buds. Microscopic studies are able to reveal the presence of fungal structures in plant tissues; however, the identification of the pathogens by microscopy is not possible in most cases due to the lack of identifiable species-specific structures. The primary infection of E. necator can have two different origins. The infection can be due to ascospores released from overwintering cleistothecia on the bark (Gadoury and Pearson, 1986; Gadoury and Pearson, 1987; Gadoury and Pearson, 1987; Gadoury et al., 2012). According to climatic conditions and the V. vinifera cultivars, overwintering mycelium in buds can also initiate the infection very early in the growing season leading to the formation of flag shoots (Corio-Costet, 2007). To set up the control strategy, it is important to determine if flag shoots are present or not. In Switzerland, cleistothecia, which can be abundant at the end of the growing season, are responsible for primary infections the next year. However, flag shoots are rarely observed and do not play a role in the epidemiology of powdery mildew in Swiss vineyards. We investigated if this correlated with the absence of mycelium in dormant buds of V. vinifera cv. Chasselas in Swiss conditions or if specific climatic conditions in spring or other factors are inhibiting the development of E. necator originating from the mycelium in buds. Dormant buds were collected in January in a non-treated plot of V. vinifera cv. Chasselas, which was naturally strongly infected with both powdery and downy mildew the year before. Microscopic analysis revealed the presence of numerous fungal structures including hyphae, spores and propagules in the fuzz of the bud (Figure 3). Even though fungal structures could be recognised, neither optical nor electronic transmission microscopy allowed for the identification of the fungi. Therefore, direct PCR was used to detect the presence of E. necator, as well as P. viticola and B. cinerea. From the 120 buds analysed, E. necator was detected by PCR in 35 buds (29%), B. cinerea in 78 buds (65%) and P. viticola could not be detected (0%). B. cinerea and E. necator were present together in 20 buds (17%). Despite the fact that flag shoots are not observed on Chasselas in Switzerland, one third of the buds in the non-treated studied plots are colonised by E. necator. These results are consistent with previous work (Rumbolz and Gubler, 2005) on V. vinifera cv. Carignane in California (USA), which reported 32.3% of buds were infected. The presence of fungus in the dormant buds does not seem to play a role in the epidemiology of the fungus in the Swiss vineyard. It could be interesting to investigate whether E. necator is present at a similar level in buds of a standard treated plot. The direct PCR method is a precious tool to demonstrate that even if flag shoots are not observed in Switzerland, E. necator is present in dormant buds. The high incidence of B. cinerea was confirmed by traditional isolation on potato dextrose agar medium, showing that B. cinerea was isolated successfully from 40 out of 120 buds analysed (33%), which represents approximately half of the incidence detected by direct PCR (65%), but this fungus did not grow on artificial media. This result shows that direct PCR is highly sensitive, perhaps because non-viable fungal propagules or conidia are detected by direct PCR. As previously reported, B. cinerea is ubiquitous; thus, it is not surprising to detect it in buds. Even if B. cinerea is not infecting young shoots under natural conditions, this may be a source of inoculum for later infections, such as bloom infection and latency (Keller et al., 2003). P. viticola, a highly specific biotrophic organism, is not able to colonise and survive in buds, which is consistent with previous reports on downy mildew epidemiology (Gessler et al., 2011).

Figure 3. Ultrastructural observations of dormant buds of Vitis vinifera cv. Chasselas. A. Global view on a longitudinal semi-thin section. B. Detailed view of A showing fungal hyphae and propagules in the fuzz (red arrows). C and D. Fungal propagules observed on a thin section. E. Fungal hyphae observed on a thin section. c: cataphyll, f: fuzz.

The direct PCR method was used for the early detection of downy mildew. Forecasting models are usually used to assess the right timing of fungicide treatments (Dubuis et al., 2012). However, it is difficult to accurately evaluate the intensity and severity of forecasted infection episodes. Based on leaf wetness duration and temperature, forecasting models can be used to precisely evaluate if the weather conditions can lead to low, middle or strong infections. However, the real severity of the infection depends on the presence of inoculum, which can be difficult to evaluate. Direct PCR on randomly sampled whole leaves could be a simple method to evaluate the level of downy mildew infection before the appearance of oil spots. Such results could help growers make decisions about spraying a curative fungicide in the three days following the infection. For example, on the 41 leaves of V. vinifera cv. Cabernet franc sampled on 17 July 2013 after a few days with strong dew in the morning, eight leaves were positive (19.5%). This is consistent with the percentage of 15% of infected leaves monitored on 5 August 2013.

Another interesting epidemiological open question in which direct PCR technique can be used is the route of infection of P. viticola that leads to grey and brown rot in grape clusters. Previous works have shown that no artificial infections were possible after BBCH 69 in the conditions of the experiment (Gindro et al., 2012). The absence of functional stomata on berries was described to be one possible cause. In the case of later downy mildew symptoms, one hypothesis discussed in this paper is that P. viticola develops systematically through the green tissues from infected leaves, tendrils, peduncles, rachis, pedicels or shoots, as previously postulated by Gregory (Gregory, 1915). Microscopic analysis of sections along a rachis, at three different points showing strong, weak and no macroscopic symptoms of downy mildew infection are shown in Figure 4. The upper part of the rachis is strongly colonised by P. viticola, and numerous hyphae and haustoria can be observed (Figure 4 B and C). In the middle part showing weak symptoms, P. viticola can be observed but with only few hyphae and haustoria (Figure 4 D and E). In the lower part of the rachis, no symptoms are observed, and P. viticola was not observed (Figure 4 F and G). Direct PCR confirmed the presence of downy mildew in the two symptomatic locations and its absence in the healthy area. This has confirmed that the hyphae observed are those of P. viticola. These observations also confirm that P. viticola is able to grow in the rachis at stage BBCH 73 and can be detected by direct PCR. These are only preliminary results, and the hypothesis that P. viticola grows systematically through the rachis to infect berries causing brown rot is under further investigation.

Figure 4. Microscopic analysis of semi-thin sections at three different points along a rachis of Vitis vinifera cv. Chasselas showing strong, weak and no symptoms of downy mildew infection. A. Grape cluster used for the study; brown rot symptoms can be observed along the upper part of the rachis. B. Semi-thin section strongly colonised by P. viticola with numerous hyphae and haustoria (red arrows). C. Detailed view of B. D. Semi-thin section with weak colonisation by P. viticola with only a few hyphae and haustoria (red arrows). E. Detailed view of D. F. Semi-thin section with no colonisation of P. viticola. G. Detailed view of F.


Direct PCR without DNA purification is a rapid method to detect and identify the presence of fungal pathogens in grapevine green and lignified tissues. This method can be used for various studies on fungal epidemiology, especially for obligate biotrophs and in field samples. The combination of microscopy and direct PCR enables the localisation and identification of the organisms present in symptomatic or non-symptomatic green tissues as well as in dormant buds. However, the method needs to be optimised for each use, especially in the removal of potential inhibitors of PCR reactions. Four examples of the usefulness of this technique in tackling different experimental hypotheses are presented. E. necator and B. cinerea are present in dormant buds, downy mildew could be detected in entire leaves shortly after rain or dew events, the presence of P. viticola in the rachis was detected and the progression of the disease could be followed during the season to better understand the appearance of brown rot. Moreover, the direct PCR method could be extended to study other pathosystems.

Acknowledgements: The authors thank Mr. Eric Remolif for production of the grapevine cuttings and helpful technical assistance, as well as Sébastien Vergne and Kees Van Leeuwen (Château Cheval blanc, Bordeaux, France) for collaborating and providing the plant material for analysis. We gratefully acknowledge the nine “Premiers Grands Crus” of Bordeaux for support to Dr. Sylvain Schnee: Château Ausone, Château Cheval blanc, Château Haut-Brion, Château Lafitte Rothschild, Château Latour, Château Margaux, Château Mouton Rothschild, Château Petrus and Château d’Yquem.


  • Aufauvre-Brown A., Tang C. M. and Holden D. W., 1993. Detection of gene-disruption events in Aspergillus transformants by polymerase chain-reaction direct from conidiospores. Current Genetics, 24, 177-178. doi:10.1007/BF00324683
  • Berthomieu P. and Meyer C., 1991. Direct amplification of plant genomic DNA from leaf and root pieces using PCR. Plant Molecular Biology, 17, 555-557. doi:10.1007/BF00040656
  • Carisse O., Tremblay D. M., Levesque C. A., Gindro K., Ward P. and Houde A., 2009. Development of a TaqMan Real-Time PCR Assay for Quantification of Airborne Conidia of Botrytis squamosa and Management of Botrytis Leaf Blight of Onion. Phytopathology, 99, 1273-1280. doi:10.1094/PHYTO-99-11-1273
  • Corio-Costet M. F. (2007). Erysiphe necator, Tec/Doc Lavoisier.
  • Dewey F. M. and Yohalem D., 2004. Detection, quantification and immunolocalisation of Botrytis species. Botrytis: Biology, Pathology and Control, 181-194.
  • Dubuis P. H., Viret O., Bloesch B., Fabre A. L., Naef A., Bleyer G., Kassemeyer H. H. and Krause R., 2012. Lutte contre le mildiou de la vigne avec le modèle VitiMeteo-Plasmopara. Revue Suisse de Viticulture Arboriculture Horticulture, 44, 192-198.
  • Gadoury D. M., Cadle-Davidson L., Wilcox W. F., Dry I. B., Seem R. C. and Milgroom M. G., 2012. Grapevine powdery mildew (Erysiphe necator): a fascinating system for the study of the biology, ecology and epidemiology of an obligate biotroph. Molecular Plant Pathology, 13, 1-16. doi:10.1111/j.1364-3703.2011.00728.x
  • Gadoury D. M. and Pearson R. C., 1986. A mechanism for ascospore discharge in Uncinula necator. Phytopathology, 76, 1145-1145.
  • Gadoury D. M. and Pearson R. C., 1987. Heterothallism and pathogenic specialization in Uncinula necator. Phytopathology, 77, 1614-1614.
  • Gadoury D. M. and Pearson R. C., 1987. Initiation and development of cleistothecia of Uncinula necator. Phytopathology, 77, 117-117.
  • Gessler C., Pertot I. and Perazzolli M., 2011. Plasmopara viticola: a review of knowledge on downy mildew of grapevine and effective disease management. Phytopathologia Mediterranea, 50, 3-44.
  • Gindro K., Alonso-Villaverde V., Voinesco F., Spring J.-L., Viret O. and Dubuis P.-H., 2012. Susceptibility to downy mildew in grape clusters: New microscopical and biochemical insights. Plant Physiology & Biochemistry (Paris), 52, 140-146. doi:10.1016/j.plaphy.2011.12.009
  • Gindro K., Pezet R. and Viret O., 2003. Histological study of the responses of two Vitis vinifera cultivars (resistant and susceptible) to Plasmopara viticola infections. Plant Physiology and Biochemistry, 41, 846-853. doi:10.1016/S0981-9428(03)00124-4
  • Gindro K., Pezet R., Viret O. and Richter H., 2005. Development of a rapid and highly sensitive direct-PCR assay to detect a single conidium of Botrytis cinerea Pers.: Fr in vitro and quiescent forms in planta. Vitis, 44, 139-142.
  • Gregory C. T. (1915). Studies on Plasmopara viticola (downy mildew of grapes). 1st International Congress of Viticulture, San Francisco, California, Geikie Press.
  • Gussow D. and Clackson T., 1989. Direct clone characterization from plaques and colonies by the polymerase chain-reaction. Nucleic Acids Research, 17, 4000-4000. doi:10.1093/nar/17.10.4000
  • Hack H., Bleiholder H., Buhr L., Meier U., Schnock-Fricke U., Weber E. and Witzenberger A., 1992. Einheitliche Codierung der phänologischen Entwicklungsstadien mono- und dikotyler Pflanzen – Erweiterte BBCH-Skala, Allgemein. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes, 44, 265-270.
  • Keller M., Viret O. and Cole F. M., 2003. Botrytis cinerea infection in grape flowers: Defense reaction, latency, and disease expression. Phytopathology, 93, 316-322. doi:10.1094/PHYTO.2003.93.3.316
  • Lepage P., Leclerc M. C., Joossens M., Mondot S., Blottiere H. M., Raes J., Ehrlich D. and Dore J., 2013. A metagenomic insight into our gut's microbiome. Gut, 62, 146-158. doi:10.1136/gutjnl-2011-301805
  • McGrath K. C., Mondav R., Sintrajaya R., Slattery B., Schmidt S. and Schenk P. M., 2010. Development of an Environmental Functional Gene Microarray for Soil Microbial Communities. Applied and Environmental Microbiology, 76, 7161-7170. doi:10.1128/AEM.03108-09
  • Miller R. R., Montoya V., Gardy J. L., Patrick D. M. and Tang P., 2013. Metagenomics for pathogen detection in public health. Genome Medicine, 5. doi:10.1186/gm485
  • Moreira D., 1998. Efficient removal of PCR inhibitors using agarose-embedded DNA preparations. Nucleic Acids Research, 26, 3309-3310. doi:10.1093/nar/26.13.3309
  • Olive D. M., 1989. Detection of entero-toxigenic Escherichia coli after polymerase chain-reaction amplification with a thermostable DNA-polymerase. Journal of Clinical Microbiology, 27, 261-265.
  • Petrosino J. F., Highlander S., Luna R. A., Gibbs R. A. and Versalovic J., 2009. Metagenomic Pyrosequencing and Microbial Identification. Clinical Chemistry, 55, 856-866. doi:10.1373/clinchem.2008.107565
  • Reynolds E. S., 1963. The use of lead citrate at high ph as an electron-opaque stain in electron microscopy. Journal of Cell Biology, 17, 208-209. doi:10.1083/jcb.17.1.208
  • Roland J. and Vian B. (1991). General preparation and staining of thin sections. Electron Microscopy of Plant Cells. J. Hall and C. Hawes. London, Academic Press: 1-66.
  • Ross-Davis A. L., Stewart J. E., Shaw J. D., Kim M. S. and Klopfenstein N. B., 2013. Metagenomic approaches for surveying forest soil microbial communities on permanent plots. Phytopathology, 103, 123-123.
  • Rossen L., Norskov P., Holmstrom K. and Rasmussen O. F., 1992. Inhibition of PCR by components of food samples, microbial diagnostic assays and DNA-extraction solutions. International Journal of Food Microbiology, 17, 37-45. doi:10.1016/0168-1605(92)90017-W
  • Rumbolz J. and Gubler W. D., 2005. Susceptibility of grapevine buds to infection by powdery mildew Erysiphe necator. Plant Pathology, 54, 535-548. doi:10.1111/j.1365-3059.2005.01212.x
  • Sachse K., 2004. Specificity and performance of PCR detection assays for microbial pathogens. Molecular Biotechnology, 26, 61-79. doi:10.1385/MB:26:1:61
  • Schnee S., Viret O. and Gindro K., 2008. Role of stilbenes in the resistance of grapevine to powdery mildew. Physiological and Molecular Plant Pathology, 72, 128-133. doi:10.1016/j.pmpp.2008.07.002
  • Segata N., Boernigen D., Tickle T. L., Morgan X. C., Garrett W. S. and Huttenhower C., 2013. Computational meta'omics for microbial community studies. Molecular Systems Biology, 9.
  • Stummer B. E., Zanker T., Harvey P. R. and Scott E. S., 2006. Detection and quantification of Erysiphe necator DNA in wine grapes and resultant must and juice. Mycological Research, 110, 1184-1192. doi:10.1016/j.mycres.2006.07.008
  • Taiz L. and Zeiger E. (2010). Plant Physiology.
  • Thiessen L. D., Mahaffee W., Keune J. A. and Grove G., 2013. Real-time detection of airborne Erysiphe necator (grape powdery mildew) inoculum with loop-mediated isothermal amplification (LAMP). Phytopathology, 103, 144-144.
  • Valsesia G., Gobbin D., Patocchi A., Vecchione A., Pertot I. and Gessler C., 2005. Development of a high-throughput method for quantification of Plasmopara viticola DNA in grapevine leaves by means of quantitative real-time polymerase chain reaction. Phytopathology, 95, 672-678. doi:10.1094/PHYTO-95-0672
  • van Belkum A., Hermans P. W. M., Licciardello L., Stefani S., Grubb W., van Leeuwen W. and Goessens W. H. F., 1998. Polymerase chain reaction-mediated typing of microorganisms: Tracking dissemination of genes and genomes. Electrophoresis, 19, 602-607. doi:10.1002/elps.1150190424
  • Vaneechoutte M., 1996. DNA fingerprinting techniques for microorganisms - A proposal for classification and nomenclature. Molecular Biotechnology, 6, 115-142. doi:10.1007/BF02740768
  • Viret O., Keller M., Jaudzems V. G. and Cole F. M., 2004. Botrytis cinerea infection of grape flowers: Light and electron microscopical studies of infection sites. Phytopathology, 94, 850-857. doi:10.1094/PHYTO.2004.94.8.850
  • Wilson I. G., 1997. Inhibition and facilitation of nucleic acid amplification. Applied and Environmental Microbiology, 63, 3741-3751.


Katia Gindro


Affiliation : Agroscope, Institut des sciences en production végétale IPV, Route de Duillier 50, 1260 Nyon, Switzerland

Nicole Lecoultre

Affiliation : Agroscope, Institut des sciences en production végétale (IPV), Route de Duillier 50, P.O. Box 1012, CH-1260 Nyon, Switzerland

Luca Molino

Affiliation : Agroscope, Institut des sciences en production végétale (IPV), Route de Duillier 50, P.O. Box 1012, CH-1260 Nyon, Switzerland

Jean-Pierre de Joffrey

Affiliation : Agroscope, Institut des sciences en production végétale (IPV), Route de Duillier 50, P.O. Box 1012, CH-1260 Nyon, Switzerland

Sylvain Schnee

Affiliation : Agroscope, Institut des sciences en production végétale (IPV), Route de Duillier 50, P.O. Box 1012, CH-1260 Nyon, Switzerland

Francine Voinesco

Affiliation : Agroscope, Institut des sciences en production végétale IPV, Route de Duillier 50, P.O. Box 1012, 1260 Nyon 1, Switzerland

Virginia Alonso-Villaverde

Affiliation : Misiôn Biolôgica de Galicia (CSIC), P.O. Box 28, 36080 Pontevedra, Spain

Olivier Viret

Affiliation : Agroscope, Institut des sciences en production végétale IPV, Route de Duillier 50, 1260 Nyon, Switzerland

Pierre-Henri Dubuis

Affiliation : Agroscope, Institut des sciences en production végétale (IPV), Route de Duillier 50, P.O. Box 1012, CH-1260 Nyon, Switzerland


No supporting information for this article

Article statistics

Views: 2183


PDF: 493