Dickeya species: an emerging problem for potato production in Europe


E-mail: john.elphinstone@fera.gsi.gov.uk


Dickeya species (formerly Erwinia chrysanthemi) cause diseases on numerous crop and ornamental plants world-wide. Dickeya spp. (probably D. dianthicola) were first reported on potato in the Netherlands in the 1970s and have since been detected in many other European countries. However, since 2004–5 a new pathogen, with the proposed name ‘D. solani’, has been spreading across Europe via trade in seed tubers and is causing increasing economic losses. Although disease symptoms are often indistinguishable from those of the more established blackleg pathogen Pectobacterium spp., Dickeya spp. can initiate disease from lower inoculum levels, have a greater ability to spread through the plant’s vascular tissue, are considerably more aggressive, and have higher optimal temperatures for disease development (the latter potentially leading to increased disease problems as Europe’s climate warms). However, they also appear to be less hardy than Pectobacterium spp. in soil and other environments outside the plant. Scotland is currently the only country in Europe to enforce zero tolerance for Dickeya spp. in its potato crop in an attempt to keep its seed tuber industry free from disease. However, there are a number of other ways to control the disease, including seed tuber certification, on-farm methods and the use of diagnostics. For diagnostics, new genomics-based approaches are now being employed to develop D. dianthicola- and ‘D. solani’-specific PCR-based tests for rapid detection and identification. It is hoped that these diagnostics, together with other aspects of ongoing research, will provide invaluable tools and information for controlling this serious threat to potato production.


The bacterial family Enterobacteriaceae includes multiple animal and plant pathogens, with the latter belonging to the genera Brenneria, Dickeya, Enterobacter, Erwinia, Pantoea and Pectobacterium (Hauben et al., 1998; Samson et al., 2005). Members of the genus Dickeya, previously known as Erwinia chrysanthemi, affect a wide range of plant hosts worldwide, particularly banana, Chrysanthemum spp., Dianthus spp., maize, potato and tomato (Burkholder et al., 1953; Samson et al., 2005). The first disease report on potato in Europe, originally identified as E. chrysanthemi and most probably belonging to the newly classified Dickeya dianthicola, was over 40 years ago (Maas Geesteranus, 1972). In most European countries, losses attributable to this pathogen have remained generally low and sporadic in the interim period, except in Switzerland where Dickeya spp. were described as predominant as far back as 1992 (Cazelles & Schwarzel, 1992). During this period, up to 25% of the potato blackleg incidences in the Netherlands (E. de Haan, NAK, Emmeloord, the Netherlands, personal communication), Belgium (J. van Vaerenbergh, ILVO, Merelbeke, Belgium, personal communication) and France (V. Hélias, INRA, Station de Pathologie Végétale, Centre de Rennes, Le Rheu Cedex, France, unpublished data) have been attributed to infections by Dickeya spp. In the last 5 years, however, potato losses caused by Dickeya spp. have increased significantly in certain cultivars in a number of European countries and in Israel (the latter a major importer of European potato seed tubers). This may be associated with the emergence of a new Dickeya pathogen that is most likely spread by trade in seed tubers and, potentially in future years, could have a larger impact as a consequence of climate change.

This review focuses on Dickeya spp. pathogenic to potato in Europe and Israel, and looks at the pathogens in terms of their geographical distribution, economic losses, biology, survival and dissemination, and symptom development. It also describes recent diagnostic methods for both research and commercial application, and considers the latest options for disease control.

The pathogen

In 1917, the genus Erwinia was established to encompass all members of the Enterobacteriaceae that were pathogenic to plants, including both pectolytic (e.g. Erwinia carotovora and E. chrysanthemi) and non-pectolytic (E. amylovora) species. Erwinia chrysanthemi was assigned to the genus by Burkholder et al. (1953) as a pathogen of chrysanthemum. Later studies revealed that strains of E. chrysanthemi cause disease on a wide variety of plant hosts, including 16 dicotyledonous families of plants in 11 orders and 10 monocotyledonous families in five orders (Samson et al., 2005; Ma et al., 2007). Because of the wide host range of E. chrysanthemi, Lelliott & Dickey (1984) subdivided the species into six pathovars, namely pvs chrysanthemi, dianthicola, dieffenbachia, paradisiaca, parthenii and zeae, based on host specificity. Later, Samson et al. (1987) developed a biovar system based on some key stable biochemical characteristics. Although Waldee (1945) proposed moving the pectolytic erwiniae into a new genus Pectobacterium based on their biochemistry, it was not until 1998 that new insights from 16S rDNA analysis led to regained impetus for such a move and to this name being largely accepted by the scientific community (Waldee, 1945; Hauben et al., 1998). Whilst the potato pathogens Pectobacterium carotovorum subsp. carotovorum (syn. Erwinia carotovora subsp. carotovora) and Pectobacterium atrosepticum (syn. Erwinia carotovora subsp. atroseptica) remain within this genus, further analysis of Pectobacterium chrysanthemi using 16S rDNA, DNA–DNA hybridization and biochemical characterization showed that it forms a distinct clade from the pectobacteria, and a new genus, Dickeya, was proposed (named after the eminent microbiologist Robert S. Dickey). Dickeya is currently divided into six species that correspond, to some extent, to both the pathovar and biovar classifications (Table 1) (Samson et al., 2005). However, since this study was undertaken, new Dickeya strains have been isolated that do not fall within these six species, and may therefore represent new Dickeya species. Throughout this review the Dickeya species will be given if known, otherwise they will be referred to as Dickeya spp.

Table 1.   Currently named members of the genus Dickeya, their synonyms and main hosts; adapted from Samson et al. (2005)
New nameSynonyms (including biovars)Hosts
DdianthicolaErwinia chrysanthemi biovars 1, 7 and 9Dianthus spp., potato, tomato, chicory, artichoke, Dahlia, Kalanchoe
Echrysanthemi pv. dianthicola
Pectobacterium chrysanthemi pv. dianthicola
DdadantiiEchrysanthemi biovar 3 (some strains)Pelargonium, pineapple, potato, Dianthus spp., Euphorbia, sweet potato, banana, maize, Philodendron, Saintpaulia
Pchrysanthemi biovar 3 (some strains)
DzeaeEchrysanthemi biovar 8 and other strains of biovar 3Maize, potato, pineapple, banana, tobacco, rice, Brachiaria, Chrysanthemum spp.
Pchrysanthemi biovar 8 and other strains of biovar 3
Dchrysanthemi bv. chrysanthemiEchrysanthemi biovar 5Chrysanthemum spp., potato, chicory, tomato, sunflower
Echrysanthemi pv. chrysanthemi
Pchrysanthemi pv. chrysanthemi
Dchrysanthemi bv. partheniiEchrysanthemi biovar 6Parthenium, artichoke, Philodendron
E. chrysanthemi pv. parthenii
Pchrysanthemi pv. parthenii
DparadisiacaEchrysanthemi biovar 4Banana, maize, potato
Echrysanthemi pv. paradisiaca
Brenneria paradisiaca
DdieffenbachiaeEchrysanthemi biovar 2Dieffenbachia, tomato, banana
Echrysanthemi pv. dieffenbachiae
Pchrysanthemi pv. dieffenbachiae

Dickeya species on potato

There is evidence that all currently described Dickeya spp. have been detected on a wide range of ornamentals in Europe, with the notable exception of D. paradisiaca (Janse & Ruissen, 1988; Samson et al., 2005; Parkinson et al., 2009; Sławiak et al., 2009b). However, D. dianthicola and emerging strains of biovar 3, for which the name ‘D. solani’ has been proposed, appear to be the only species that have spread to potato in Europe. Future introductions of new Dickeya spp. to potato cannot be excluded in terms of potential infection pathways via other plant species and adaptability to current and future climatic conditions. Previous reports of the isolation of E. chrysanthemi biovar 5 from potato in the Netherlands (Janse & Ruissen, 1988) and biovars 5 and 6 in Spain (Palacio-Bielsa et al., 2006) require a more complete identification of the Dickeya spp. involved. Similarly, reported findings of D. dadantii (biovar 3) and D. dieffenbachiae (biovar 2) in Germany also require further investigation, since these would be the first reports of these species on potato in Europe (Sławiak et al., 2009b).

The first European report of Dickeya spp. (E. chrysanthemi) on potato was from the Netherlands in the 1970s (Maas Geesteranus, 1972), and it has since been found in England (Parkinson et al., 2009), France (Hélias, 2006), Hungary (J. Nemeth, Plant Protection and Soil Conservation Service, Hungary, personal communication), Jersey (D. E. Stead, Fera, York, UK, personal communication), Sweden (Persson, 1991), Belgium (J. van Vaerenbergh, personal communication), Switzerland (Cazelles & Schwarzel, 1992), Poland (Sławiak et al., 2009a), Finland (Laurila et al., 2006), Scotland (Cahill et al., 2010), Spain (Palacio-Bielsa et al., 2006), Georgia (L. Tsror, ARO, Negev, Israel, unpublished data) and Israel (Lumb et al., 1986; Tsror et al., 2009) (Fig. 1). By the early 1990s, E. chrysanthemi was reported to be the most frequent bacterial pathogen on seed tubers in W. Switzerland, followed by P. atrosepticum (Cazelles & Schwarzel, 1992).

Figure 1.

 Known distribution of Dickeya dianthicola on all hosts in Europe, together with distribution of ‘Dickeya solani’ and other Dickeya species on potato in Europe. Distribution of D. dianthicola on all hosts (updated from CAB International, 2005) (•); Dickeya spp. on potato (¤); ‘D. solani’ on potato (o).

Prior to 2004, almost all European potato isolates of Dickeya tested appeared to be D. dianthicola (Parkinson et al., 2009; Sławiak et al., 2009b). This species was first detected causing stunting and slow wilting of Dianthus in the early 1950s in Denmark, the Netherlands and the UK (Hellmers, 1958). Dickeya dianthicola was subsequently listed as a quarantine organism (EPPO A-2) on Dianthus (Council Directive 2000/29/EC), but has also been found to cause disease on potato, tomato, chicory and artichoke, as well as on ornamental genera such as Begonia, Dahlia, Freesia, Hyacinthus, Iris, Kalanchoe and Zantedeschia (Samson & Nassan-Agha, 1978; Dickey, 1979; Janse & Ruissen, 1988; Lee et al., 2002; Samson et al., 2005; van Doorn et al., 2006). Bradbury (1986) recorded the presence of D. dianthicola on Dianthus in Denmark, England, France, Germany, Italy, the Netherlands, Norway, Poland, Romania, Sweden and Greece. Recent studies found that D. dianthicola strains isolated from eight host plants, almost exclusively from Europe, showed little sequence diversity despite encompassing multiple isolates recovered from Dianthus (Fig. 1) and (more recently) potato (Parkinson et al., 2009; Sławiak et al., 2009b). This observation suggests that D. dianthicola may have initially spread to potato in Europe from other host plants, and presumably from Dianthus. Dickeya dianthicola has been successfully controlled on Dianthus through strict glasshouse hygiene and certification of planting material.

Several recent studies based on sequence data, biochemistry and REP-PCR analyses of potato strains isolated after 2004 from the Netherlands, UK, Finland, Poland, Israel and elsewhere in Europe (Fig. 1) have identified a new clade of Dickeya belonging to biovar 3. This clade, identified independently as group I by Laurila et al. (2008), DUC-1 by Parkinson et al. (2009) and clade IV by Sławiak et al. (2009b), appears to be clonal between countries, is closely related to a Dutch strain from hyacinth, and is different from the six known Dickeya species (Laurila et al., 2008; Czajkowski et al., 2009b; Parkinson et al., 2009; Sławiak et al., 2009b; Tsror et al., 2009; Kowalewska et al., 2010). It may, therefore, represent a new species, for which the name ‘D. solani’ has been proposed but has not yet been formally accepted. Since 2005, ‘D. solani’ has commonly been isolated from seed potato tubers in the Netherlands, even though a wide range of different cultivars and locations have been sampled, and ‘Dsolani’ also appears to predominate on certain cultivars in Israel (Czajkowski et al., 2009a,b; Tsror et al., 2009). In Spain, a biovar 3 strain was isolated from potato in 2002, but its sequence similarity to ‘Dsolani’ has not yet been established (Palacio-Bielsa et al., 2006). However, ‘D. solani’ has been identified on potatoes exported from Spain (G. Saddler, SASA, Edinburgh, UK, unpublished data). As with D. dianthicola, strains of ‘D. solani’ may have spread to potato from other host plants since strains belonging to this genetic clade from potato are very similar (as determined by REP-PCR, 16S rDNA and dnaX sequence analyses) to strains isolated recently from hyacinth bulbs (Sławiak et al., 2009b).

A number of other, as yet unclassified, clades of Dickeya have also recently been isolated in Europe from hosts other than potato (Parkinson et al., 2009). These include the unidentified clades DUC-2, isolated mostly from a range of monocotyledonous species, and DUC-3, represented by a single strain from Aglaonema spp. DUC-3 was recently isolated from river water in Scotland, but has not yet been found on potato, although it is able to rot potato tubers (Cahill et al., 2010). Two further clades, highly distinct and probably ranking at species level, have been recognized; the first, isolated from sugarcane in Australia (SLC-1), has not been found in Europe, but the second (SLC-2), of unknown host, has so far only been isolated from river water in England and Finland (Laurila et al., 2008; Parkinson et al., 2009).

There are reports worldwide of isolates simply identified as E. chrysanthemi on many hosts (including potato), but in most cases identification to the pathovar (Lelliott & Dickey, 1984) or biovar level (Samson & Nassan-Agha, 1978) was not carried out. It is not possible, therefore, to assign a Dickeya sp. to these isolates without further sequence analysis. Outside Europe, D. dianthicola has been reported from ornamental hosts in Colombia, Japan, New Zealand and the USA (New York, Pennsylvania and Texas) (Bradbury, 1986). To date, no potato isolates from outside Europe have been confirmed as D. dianthicola, with the exception of one sample from Bangladesh that may have been grown from European seed tubers. Further testing is required to confirm whether isolates identified as ‘Echrysanthemi’ and reported on crops grown from European seed tubers in North Africa, Cuba and elsewhere, can be classified as D. dianthicola or, as in the recent cases in Israel, as ‘D. solani’ (Parkinson et al., 2009; Sławiak et al., 2009b; Tsror et al., 2009).

Dickeya spp. identified on potato appear to be distinct on different continents. These include D. chrysanthemi in the USA and Taiwan (Parkinson et al., 2009; Sławiak et al., 2009b), D. dadantii (also identified as biovar 3) in Brazil (Parkinson et al., 2009), Peru (DeLindo et al., 1978; Sławiak et al., 2009b) and Zimbabwe (Ngadze et al., 2010), and D. zeae (also identified as biovar 3) in Australia and Papua New Guinea (Cother, 1980; Cother et al., 1992; Parkinson et al., 2009; Sławiak et al., 2009b). Distinct genotypes of D. zeae were recently isolated from river water in Scotland and England, but have not yet been found on potato in Europe (Cahill et al., 2010; J. G. Elphinstone, Fera, Sand Hutton, York, YO41 1LZ. UK, unpublished data).

Economic losses caused by Dickeya species on potato in Europe

Quarantine status

EPPO currently lists E. chrysanthemi as an A2 quarantine organism of carnations and chrysanthemums, although it is specified that the quarantine pests concerned are E. chrysanthemi pvs dianthicola and chrysanthemi (OEPP/EPPO, 1982, 1988, 1990), which equate to D. dianthicola and D. chrysanthemi bv. chrysanthemi, respectively. It has, however, been proposed that they should be delisted once national nuclear stock certification schemes have been agreed upon for these ornamentals. There are no current quarantine implications in Europe for D. dianthicola or any other Dickeya species on potato, where it has widely been considered that the risk from Dickeya spp. can be adequately covered by national nuclear stock (in vitro pathogen-tested microplants) and seed tuber certification schemes. However, in Israel it is considered to be a quarantine organism (Tsror et al., 2009) and in Scotland (a community grade region within the European Union that produces approximately 99·5% of its own seed potato tubers) a unique decision was taken in 2010 to introduce zero tolerance for all Dickeya spp. on potatoes as part of its Seed Potato Classification Scheme [The Seed Potatoes (Scotland) Amendment Regulations 2010].

Potential economic impact

In Israel, yield reductions of 20–25% resulting from Dickeya infections have been recorded on various potato cultivars, where disease incidence was greater than 15% (Tsror et al., 2009). In Finland, in a 1-year field trial, a comparison was made between direct losses caused by D. dianthicola and ‘D. solani’ measured by tuber and stem rot (Laurila et al., 2008). On average, no difference was found in the percentages of tuber decay (5–6%) between these pathogens but the percentage of stem rot was much higher for D. dianthicola (73% vs. 20%). However, most direct losses to potato production in Europe caused by Dickeya have occurred as a result of downgrading or rejection of potatoes during seed tuber certification. Since national certification tolerances differ, the economic impact varies from country to country. Strict tolerances in the Netherlands have led to increased direct losses of up to €30M annually (Prins & Breukers, 2008) as a result of downgrading and rejection of seed tuber stocks caused by blackleg. However, it is not possible to differentiate losses caused by Pectobacterium and Dickeya. The appearance of symptoms during seed tuber certification and growing-crop inspections depends largely on the prevailing climate, especially during the early part of the season. The inoculum level on seed and the cultivar used may also play a role in symptom development, although inoculation of tubers with as little as 40 cells per gram of potato peel was sufficient to cause 30% and 15% diseased plants in field experiments in the Netherlands in 2005 and 2006, respectively (van der Wolf et al., 2007). During official inspections in England and Wales, some potato crops found to be affected by either D. dianthicola or ‘D. solani’ showed foliar symptoms ranging from <1% to 30% (J. Elphinstone, Fera, Sand Hutton, York, YO41 1LZ. UK, unpublished data).

Crop losses can occur for seed tuber growers, suppliers and exporters if these tubers are exported to warmer climates and break down as a result of the presence of Dickeya spp. With the emergence and spread of ‘D. solani’, together with the effects of climate change, more frequent losses of this kind can be expected. Data from Israel indicate the scale of potential losses that can be expected when seed tubers latently infected with D. dianthicola or ‘D. solani’, are grown in warmer climates (Lumb et al., 1986; Tsror et al., 2006). For example, in spring 2005, using seed tubers imported to Israel from Europe, a severe outbreak of disease (subsequently found to be caused by ‘D. solani’) was observed in more than 200 ha across different locations. Five cultivars were involved, with disease incidence ranging from 5% to 30% (8·2% average). In addition to foliar wilting symptoms, rotted progeny tubers were also observed in the field. In one cultivar, a high wilt incidence (30%) was observed and, when visually healthy progeny tubers were replanted, a further wilt incidence of 10–15% was observed in the following autumn–winter season. In spring 2006, the disease was again observed in more than 260 ha in seven cultivars, with disease incidence ranging from 2% to 30% (10% average) (Tsror et al., 2009). In spring 2009, 12 cultivars were involved with disease incidence of 0·5–30% (L. Tsror, unpublished data). When healthy potato seed tubers were planted in two locations in 2005, following the original diseased crop, no transmission of the disease to the healthy crop was observed and the pathogen was not detected on the progeny tubers. It was thus concluded that the pathogen is not significantly soilborne and its primary means of dispersal is in seed tubers.

Biology, dissemination and survival of the pathogen

Factors influencing disease development

Factors favouring disease development on potato caused by Dickeya spp. are, on the whole, similar to those for P. atrosepticum and include damage and lack of cleanliness at grading, poor soil drainage, presence and increasing level of the pathogen on seed tubers, over-irrigation, wet spring weather, damage at harvest, and lack of adequate ventilation at storage. However, there are factors that may influence disease development differently between Dickeya spp. and P. atrosepticum (or even between Dickeya spp.). These include inoculum level, cultivar susceptibility, speed of migration through the plants’ vascular system, temperature and relative aggressiveness. Unfortunately, there is little published information on many of these factors and their differential roles in disease promotion are largely unknown. However, information that is available is outlined below.

Lower inoculum densities of Dickeya spp. than of P. atrosepticum are required for disease development (D. E. Stead, personal communication; I. Toth, SCRI, Invergowrie, Dundee, DD5 2DA, UK, unpublished data; van der Wolf et al., 2007). In one study, all or most susceptible tubers inoculated with 10 cells of ‘D. solani’ and strains of D. dianthicola (with higher optimal temperatures), developed disease at both 21 and 27°C. This was compared to a much lower proportion of tubers that developed disease when inoculated with the same level of P. atrosepticum and D. dianthicola (with lower optimal temperatures) at 21°C, with no disease development at 27°C (I. Toth, unpublished data). To support these findings, ‘D. solani’ strains were shown to cause more severe losses than D. dianthicola, P. atrosepticum and P. carotovorum subsp. carotovorum (Lojkowska et al., 2010). However, no difference in average incidences between ‘D. solani’ and D. dianthicola were found in a 3-year field study in the Netherlands (Czajkowski et al., 2010). The aggressiveness of 40 ‘D. solani’ strains evaluated in a tuber maceration bioassay (incubation for 48 h at 30°C) indicated a considerable variability among the tested strains, ranging from 0·4 to 4·0 g of macerated tissue (L. Tsror, unpublished data).

There is currently little data available on the susceptibility of cultivars to Dickeya spp. It is mostly assumed, possibly wrongly, that susceptibility ratings for P. atrosepticum are suitable for assessing potential resistance to Dickeya spp. However, preliminary work in England (D. E. Stead, personal communication) has shown that all major potato cultivars tested were susceptible to D. dianthicola, with some variation in symptom severity following inoculation in small-scale field trials. Certain cultivars have also been identified as highly susceptible to Dickeya spp. under Israeli conditions and, as a consequence, are now being imported to a much lesser extent (L. Tsror, unpublished data).

Dickeya solani’ is able to colonize roots of potato plants from soil within 1 day, irrespective of root damage (Czajkowski et al., 2010). In this study, the pathogen was found in stolons and stems 15 days after soil inoculation. Although no comparative study was done, there is circumstantial evidence that Dickeya spp. are better invaders of vascular tissue of potato plants than P. atrosepticum. Recent unpublished data has shown that there is no obvious difference in the rate of mother tuber breakdown between the two pathogens, although the effect of temperature on such breakdown has yet to be assessed. Detailed studies showed that, like Dickeya spp., P. carotovorum subsp. carotovorum may also be found at high concentrations in the stolon ends of tubers, whereas for both pathogens relatively low densities were evident in the peel and more deeply located tuber tissue (Czajkowski et al., 2009a). No cultivar effects were evident nor was there any significant difference in the distribution of either pathogen.

Temperature is perhaps the most important factor in determining whether disease in any one season will be predominantly caused by Dickeya spp. or P. atrosepticum, and there are both published and unpublished data to support this. Lumb et al. (1986) found that in Israel symptoms caused by Dickeya spp. tended to develop when temperatures exceeded 25°C, whilst P. atrosepticum predominated below 25°C. Pérombelon & Hyman (1986) also showed that temperature plays a vital role in determining which pathogen predominates in causing disease symptoms during and between growing seasons. Similarly, investigations in the Netherlands suggest a correlation between spring/summer temperatures and the genus responsible for causing the symptoms in the field (J. van der Wolf, PRI, NL-6700 AB, Wageningen, the Netherlands, unpublished data). However, in all these cases, the Dickeya species involved was not determined.

The relative aggressiveness of different isolates is closely linked to temperature, although pathogen testing in any particular study is often performed at only a single temperature, making comparisons between studies and, therefore, isolates difficult. For example, studies in Israel used 28–30/22–24°C day/night temperatures (Tsror et al., 2009), compared to 28/18°C (day/night temperatures) in Spanish investigations (Palacio-Bielsa et al., 2006), and 22°C (for tubers) and 23/21°C (day/night temperatures for stems) in Finland (Laurila et al., 2008). However, current evidence suggests that ‘D. solani’ is able to grow at temperatures as high as 39°C, and is more aggressive than D. dianthicola at higher temperatures, as determined by greenhouse studies and experiments in high-temperature regions in Israel (Laurila et al., 2008; Sławiak et al., 2009b; Tsror et al., 2009). The opposite may be true at lower temperatures, where field experiments in Finland showed that, on average, D. dianthicola resulted in the highest incidence of diseased plants (Laurila et al., 2008). In that study, isolates of ‘D. solani’ were found to be most variable in aggressiveness when tested in vitro. It was found that both ‘D. solani’ strains and a P. atrosepticum control strain were more aggressive than isolates of D. dianthicola on tubers and stems at 22–23°C (Laurila et al., 2008). A ‘D. solani’ isolate from Israel was highly aggressive in potato stem assays at 28–30°C daytime temperature, whilst P. atrosepticum failed to cause disease under these conditions (Tsror et al., 2009). Similarly, a Dickeya biovar 3 isolate from Spain was more aggressive than a biovar 1 isolate in in vitro stem tests at 28°C, with no obvious difference in symptoms between cultivars used (Palacio-Bielsa et al., 2006). Recent work showed that D. dianthicola isolates also exhibit variable aggressiveness, which manifests at different optimal temperatures between 21 and 27°C. The least aggressive of these were similar in aggressiveness to isolates of P. atrosepticum on tubers, and had similar optimal temperatures of around 21°C. However, at this temperature, and up to 27°C, aggressiveness on stems appeared to be greater for D. dianthicola isolates than for P. atrosepticum. Other, more aggressive D. dianthicola isolates had a level of aggressiveness significantly higher than P. atrosepticum on both stems and tubers at the higher optimal temperatures (27°C), similar to or less than that of a single test isolate of ‘D. solani’ (IPO2222) at the same temperature (I. Toth, unpublished data).

Risk of spread via latently infected propagation material

Dickeya spp. spread over long distances, and across national borders, in infected vegetative propagating material. The most important means of dissemination for bacterial pathogens of potato is the movement of latently infected seed tubers (Pérombelon & Kelman, 1980; Tsror et al., 1999, 2009). The pathogen can be carried on the tuber surface and in lenticels (as for Pectobacterium spp.), but is also likely to be found in the tuber vascular system, which it enters systemically via the stolon from the infected mother plant or via root infection (Underberg, 1992; Czajkowski et al., 2009a, 2010).

Risk of spread in watercourses

There have been several reports of E. chrysanthemi detection in surface water, although the corresponding Dickeya spp. have yet to be determined in most cases. Cother & Gilbert (1990) detected E. chrysanthemi in two major river systems in Australia. The pathogen was more readily isolated when stream weeds and sediment were included in the water sample and only when the temperature was 16°C or higher. In sterile water, the bacterium was shown to survive for at least 211 days at 16°C (van Doorn et al., 2008). van Doorn et al. (2008) found large differences in survival rates of E. chrysanthemi in sterile water, ranging from 7 days in tap water, 21 days in ditch water, 49 days in phosphate buffered saline to over 154 days in water sampled from a rain water basin. It seems that both the buffering of water and the presence of nutrients favour survival. However, survival of E. chrysanthemi in water was found to be shorter than for Pectobacterium spp. Olsson (1985), reported the isolation in 1976 of E. chrysanthemi from Solanum dulcamara growing in a watercourse used for irrigation in Sweden, and in 1983 the pathogen was isolated directly from water samples obtained from the same source. The bacteria were shown to infect potatoes after artificial inoculation and to be transmitted to subsequent generations. Dickeya has also been isolated from irrigation water sources in the Netherlands (Van Vuurde & de Vries, 1992). The pathogen did not survive during the winter and was only detected if the water temperature was above 10°C. This water contamination appeared not to be related to the presence of potato fields. Isolates were found to belong to biovar 3, but the bacteria were not identified to species level.

Other studies in the Netherlands indicated that in autumn a high percentage of the surface water was contaminated with both P. atrosepticum and Dickeya spp., with densities between 100 and 1000 cells L−1 (Roozen, 1990). A similar observation was recently made in Finland (Laurila et al., 2006, 2008, 2010), where some of the soft rotting bacteria isolated from river water samples were found to be identical to ‘D. solani’. ‘Dickeya solani’ was also recently detected in river water in SE Scotland, but not in Scottish seed potato tubers (Cahill et al., 2010). In addition, DUC-3 (defined by Parkinson et al., 2009 and as yet unnamed) and D. zeae, previously found on a number of exotic ornamentals, were also found in Scottish rivers. In each case, the population densities remained constant after repeated annual sampling, and there was no evidence of other Dickeya spp. being present (Cahill et al., 2010). Isolates from water samples taken from a number of unconnected rivers in SE England were found to contain either a single sequence type of Dzeae (in seven different catchments) or a single sequence type of a non-identified Dickeya spp. (in three different catchments) (N. Parkinson, Fera, York, UK, personal communication). Interestingly, this latter Dickeya spp. was also found in river water in Finland (Laurila et al., 2008, 2010), but did not cause stem rotting after inoculation, although it was pectolytic on potato tuber tissue.

In Europe, there has been little or no correlation between Dickeya spp. isolated from river water and those found on potato. In Australia, however, irrigation water was found to be the likely source of infection of potatoes (Cother et al., 1992). Two different Dickeya spp. were isolated from the headwaters of the Murrumbidgee River and the source of the Murray River in New South Wales. Isolates from the latter have since been confirmed as D. zeae (Parkinson et al., 2009). In both cases, biochemical, fatty acid and DNA characterization methods showed that isolates from potatoes were identical to those from the river water with which they were irrigated. In Florida, high populations of Dickeya spp. able to infect Dieffenbachia spp. were also detected in irrigation ponds containing recycled water (Norman et al., 2003).

Survival in soil

Survival studies reported in the scientific literature often fail to specify the Dickeya spp. involved, and results are thus difficult to extrapolate to European conditions. However, in general, it appears unlikely that the pathogen can overwinter freely in soils. A maximum survival period of 12 months for E. chrysanthemi in soil was reported for potting media in glasshouses (Haygood et al., 1982), whilst studies in Italy showed that Dickeya isolates from Dianthus cannot survive in plant-free soil for more than 6 months (Garibaldi, 1972). In plant-free soils in tropical areas, survival periods for E. chrysanthemi were found to vary between 7 days (Lim, 1975) and 3 months (Rangarajan & Chakravarti, 1970), and to be dependent on temperature, moisture level and pH (Anilkumar & Chakravarti, 1970). Erwinia chrysanthemi was found to survive in soil for up to 38 days at 8°C, 22 days at 20°C and 12 days at 30°C: at low soil moisture levels (30%) longer survival periods were found than at higher soil moisture levels (>60%). Erwinia chrysanthemi was also shown to survive for twice as long in sterilized soil than in unsterilized soil, demonstrating the antagonistic role of the soil microflora on the survival of E. chrysanthemi (Anilkumar & Chakravarti, 1970; Rangarajan & Chakravarti, 1970; Garibaldi, 1972). Moreover, D. dianthicola and ‘D. solani’ seeded to soil persisted only for a maximum of 3 weeks, irrespective of soil type, temperature and humidity (van der Wolf et al., 2007). However, persistence of Dickeya spp. for longer periods in association with crop residues in soil cannot be excluded. In the Netherlands, infected crop residues buried in soil in the autumn showed some transmission to Dickeya-free mini-tubers in the next growing season (Velvis & van der Wolf, 2008). Other recent studies in the Netherlands suggest that Dickeya isolates from potato and hyacinth cannot survive for more than 7 days when added to different soils at 6°C and 50% field moisture capacity (compared with 942 days for Pectobacterium isolates) (van der Wolf et al., 2009). Nevertheless, potato crops multiplied twice in the field from pathogen-free mini-tubers were observed to have 20–56% infection by Dickeya spp. in the harvested tubers. Confirmation of such high primary infection rates suggests that the pathogen is transferred via contaminated farm machinery, can survive in the potato-growing environment (e.g. on plant debris or on alternative hosts, either other crops or weed species) and/or is transmitted from outside of the cropping environment (e.g. via irrigation water, aerosols or insects).

With the exception of a short report about the infection of Solanum dulcamara in Sweden (Olsson, 1985), there was until recently little information on the potential for survival of D. dianthicola and other Dickeya spp. in weed hosts. However, given the high host diversity across and sometimes within the Dickeya spp., it is highly likely that wild host plants could play a role in the survival and spread of the pathogen. To study dissemination to weeds, surveys were conducted in potato fields in Israel, where Dickeya-infected potato plants were detected during two consecutive spring seasons (2009 and 2010). Symptomless plants of 12 species of local weeds were randomly collected: Cyperus rotundus, Orobanche aegyptiaca, Amaranthus spinosus, Polygonum equisetiforme, Chenopodium sp., Heliotropium sp., Centaurea iberica, Sorghum halepense, Malva nicaeensis, Cynodon dactylon, Amaranthus blitum and Solanum elaeagnifolium. Dickeya spp. were isolated in both years but only from the perennial weed C. rotundus, with 6·7% (2009) and 14·3% (2010) of these plants harbouring the pathogen (Tsror et al., 2010), suggesting that there may be some host specificity.


Distinction from symptoms caused by P. atrosepticum and P. carotovorum

Prior to the reclassification of E. chrysanthemi into species within the genus Dickeya it was unclear whether symptoms caused by Dickeya spp. on potato in Europe were similar to those of P. atrosepticum or were sufficiently different to allow disease diagnosis. Recent work showed that there are at least two species of Dickeya causing this disease in Europe (D. dianthicola and ‘D. solani’), with isolates of each apparently showing large variations in pathogenicity and optimal temperature (Laurila et al., 2008; I. Toth, unpublished data). Observations made in the Netherlands suggest that the same symptoms may be caused by D. dianthicola, ‘D. solani’ or P. atrosepticum (J. M. van der Wolf, PRI, NL-6700 AB, Wageningen, the Netherlands & E. de Haan, NAK, 8300 BC, Emmeloord, the Netherlands, unpublished data), e.g. typical blackleg symptoms (Fig. 2a). However, observations from field experiments in Finland suggest that symptoms caused by Dickeya spp. differ from those of blackleg caused by P. atrosepticum (Laurila et al., 2010). It is, therefore, highly likely that there is no simple relationship between symptomatology caused by isolates of P. atrosepticum and Dickeya spp., but rather that there may be a range of symptoms depending on the species, isolate, environmental conditions and even cultivar involved. However, generally it appears that disease caused by Dickeya spp. under warm, wet conditions leads to stem rotting with symptoms similar to those of P. atrosepticum, whilst under lower humidity less rotting is observed but with wilting, increased leaf desiccation, stem browning and hollowing of the stem (Lumb et al., 1986; De Vries, 1990; Palacio-Bielsa et al., 2006; Tsror et al., 2009).

Figure 2.

 Disease symptoms in potato tubers and stems caused by Dickeya species: (a) typical blackleg symptoms caused by ‘Dickeya solani’; (b) ‘Dickeya solani’ soft rot of developing progeny tuber extending from the stolon; (c) soft rot of daughter tubers developing from the stolon; (d) initial wilt in upper leaves; (e) increased levels of necrosis in the upper leaves and wilt and desiccation in the lower leaves; (f) D. dianthicola rotting mother tuber; (g) internal stem necrosis or rotting extending from the stem base, but with the stem base appearing externally healthy; (h) ‘Dickeya solani’ on imported potato causing cheesy rot and break down of the vascular ring similar to ring rot or brown rot. (b, f) Fera Crown Copyright; (c, d, e) courtesy of L. Tsror, Gilat Research Centre, Israel; (h) courtesy of Eric Anderson, Scottish Agronomy.

Symptoms of soft rot on potato tubers, as described by Powelson & Franc (2001), appear to be similar whether caused by Dickeya or Pectobacterium spp. Tuber soft rot ranges from a slight vascular discoloration to complete decay. Affected tuber tissue is cream to tan in colour and is soft and granular. Brown to black pigments often develop at the margins of decayed tissue (Fig. 2b). Lesions usually first develop in lenticels, at the site of stolon attachment or in wounds (Fig. 2c). Recent work at SCRI showed that Dickeya spp., particularly at higher temperatures (27°C), cause more severe rots than P. atrosepticum and are more likely to produce a creamier, cheesy rot (I. Toth, unpublished data).

The foliar symptoms traditionally associated with D. dianthicola in Europe and Israel occur in warm, dry growing conditions (Lumb et al., 1986; Tsror et al., 2009). The first symptom is a wilt of the top leaves, with subsequent desiccation around the margins and eventually of the entire leaves. These symptoms become visible in the lower leaves and, in extreme cases, the whole plant or stem dries out (Fig. 2d,e). Often, only one stem per plant is affected. Symptom development is usually associated with soft rotting of the mother tuber (Fig. 2f), but the soft rot symptoms do not always extend up the stolon or stem, either externally or internally, as observed with blackleg caused by P. atrosepticum. Vascular tissues stain brown from the stem base, progressing upwards and occasionally resulting in necrosis and hollowing of the stem (Fig. 2g). Externally, the stems usually remain green until leaf desiccation is complete. Under warm, dry conditions, symptoms usually first appear when the air temperature exceeds 25°C (Lumb et al., 1986).

Other Dickeya spp. (D. dadantii and D. zeae), generally found on potato in warmer, humid tropical and subtropical environments, cause symptoms that are indistinguishable from those of blackleg disease caused by P. atrosepticum in cooler environments (Cother, 1980; DeLindo & French, 1981). Plants affected by these organisms show wilting, stunting and chlorosis, and a brown to black soft rot at the stem base, extending upwards from the rotting mother tuber, with eventual total collapse of the plant. When disease occurs before or just after emergence, missing plants (blanking) are observed in the crop. In low-temperature growing regions, infections often start with dark colouring of upper leaves, followed by chlorosis and wilting. As the disease progresses, stems wilt or show blackleg symptoms. Contamination of foliage and subsequent crop damage caused by heavy rainfall, hail, insects or human activities can result in aerial stem rot (van der Wolf & De Boer, 2007). Rather than soft rot extending from the stem base, external symptoms often appear higher up the plant. Such plants rapidly develop internal stem rotting from their base, but externally the stem base appears healthy (Fig. 2g). ‘Dickeya solani’ commonly causes rotting of developing progeny tubers in the field, especially under warm growing conditions (Fig. 2c).

Distinction from other diseases

Whilst typical blackleg and soft rot diseases caused by Pectobacterium spp. are easily distinguishable from other diseases on the basis of visual inspection, symptoms caused by D. dianthicola or ‘D. solani’ under warm, dry conditions may be confused with those of other wilting diseases. In Israel, symptoms caused by Dickeya spp. were indistinguishable from those of wilt caused by Verticillium dahliae or those caused by natural plant senescence (Lumb et al., 1986). Tuber symptoms caused by Dickeya spp. might also be confused with those of other bacterial diseases of potato, including brown rot (caused by Ralstonia solanacearum) and ring rot (caused by Clavibacter michiganensis subsp. sepedonicus), as ‘D. solani’ has been shown in some cases to produce a discoloration and/or rotting of the vascular ring within progeny tubers (Fig. 2h).


Seed certification

Pectobacterium atrosepticum and Dickeya spp. are both regarded as seed tuber pathogens, and their control within the European Community is largely through seed tuber classification in line with national and EC legislation. Most seed tubers are derived initially from pathogen-tested nuclear stock microplants, which ensures that they are initially free from bacterial pathogens (Stead, 1999). This high health status was confirmed in a 4-year survey where 25–30 seed lots of minitubers belonging to different cultivars were tested and found to be free of the potato pathogens (Velvis & van der Wolf, 2008). Multiplication of seed potato tubers over a limited number of field generations (usually three to five) therefore helps to minimize the buildup of contamination and infections (but tends to sharply increase the price of seed tubers for the end-user).

The seed tuber classification schemes set tolerances for the levels of soft rot and blackleg diseases encountered during visual inspections of growing crops and of harvested tubers after grading and dressing, and prior to marketing. For high-grade seed tubers, there is generally zero tolerance for blackleg and soft rot diseases in both field and tuber inspections. However, tolerances in the later stages of seed tuber multiplication vary between countries (Table 2). In addition, removal (roguing) of some or all blackleg plants is often allowed between inspections. Records of the levels of blackleg and soft rot encountered at such inspections therefore provide some guidance on general health across stocks and generations but do not accurately reflect the status of latent infections, the pathogen species present or the general risk of blackleg or soft rot disease development in subsequent generations. In the majority of member states of the European Union, there are no official postharvest testing programmes for Dickeya or Pectobacterium spp., although voluntary testing services offered in some countries (including the UK and the Netherlands) can provide useful decision support for growers and store managers. Scotland is unique within the EU in enforcing a system where all non-indigenous seed tubers must be tested prior to planting to ensure that they are free of Dickeya spp. (http://www.scotland.gov.uk/Topics/farmingrural/Agriculture/plant/18273/PotatoHealthControls/PotatoQuarantineDiseases/Dickeya). It has also introduced a zero tolerance for blackleg caused by Dickeya spp. in its seed tuber classification scheme, using a system based on field inspection backed by laboratory testing, in which high-risk crops (non-Scottish origin) are targeted but 10% of indigenous production is also surveyed to ensure freedom from Dickeya spp.

Table 2.   Tolerances for blackleg in European seed potato tubers
  1. NA, not applicable.

  2. aPre-basic seed tuber potatoes are field-grown potatoes produced from mini-tubers grown under controlled conditions, which were initially propagated from disease-tested microplants. There are no disease tolerances for pre-basic seed stocks, which must be completely free of disease symptoms.

  3. bBasic seed potato tubers are derived from pre-basic seed tubers. There are disease tolerances which relax with each multiplication. Pre-basic and basic seed tubers are intended for the production of seed tuber crops.

  4. cCertified seed tubers are mainly intended for the production of potatoes for consumption. Some countries in the European Union, e.g. Scotland, prohibit the planting of certified seed tubers.

  5. dNo distinction made between blackleg caused by P. atrosepticum, P. carotovorum or Dickeya spp.

  6. eDerived from data produced by United Nations Economic Commission for Europe Working Party on Agricultural Quality Standards (http://www.unece.org/trade/agr/welcome.htm)

England & Walesd00·25–1·02·0
Scotland –Pectobacterium spp.00·25–1·0NA
UK (Scotland) –Dickeya spp.00NA

On-farm measures

On-farm control measures currently recommended for Dickeya spp. are essentially the same as for P. atrosepticum (Pérombelon, 1981), largely because there is insufficient data available at present to support Dickeya-specific measures. Contaminated seed tubers, the use of contaminated machinery and equipment during cultivation, harvesting and grading, and possibly environmental contamination are considered to play important roles in the introduction of Dickeya spp. to the potato crop.

In areas where Dickeya spp. are not present, biosecurity measures should be aimed at avoiding introduction of the pathogen. Infected seed tubers represent the most likely source of introduction to a new area. Almost all new findings of D. dianthicola and ‘D. solani’ have been traced back to the seed source (Lumb et al., 1986; Laurila et al., 2006; Sławiak et al., 2009a; Tsror et al., 2009). Measures to ensure that seed tubers originate from Dickeya-free areas, crops or even stocks of cultivars require a consistent level of monitoring of crops and their environment for presence/absence of the pathogens.

One possible mechanism, which is gaining support from the potato industry in the UK, is membership of an assured seed production scheme, e.g. Safe Havens (http://www.potato.org.uk/department/export_and_seed/safe_haven/index.html?menu_pos=seed). This scheme ensures that only disease-free microplants can enter the production chain and that field-grown generations can only be grown on agricultural units that cannot handle seed tubers produced outside the scheme. In this way, healthy planting material is passing through the production chain, with no possible avenue for the introduction of infection from sources of seed tubers.

Once Dickeya has been introduced, there are a number of measures that can reduce the risk of spreading the pathogens within and between crops. Mechanical harvesting is likely to increase the opportunity for pathogen spread from tuber to tuber as well as facilitating infection through wounds. Avoidance of mechanical harvesting during the early phases of pre-basic seed tuber multiplication (more widely practised during seed production using clonal selection than in seed production using mini-tubers) may reduce the rate of contamination of healthy stocks. Cleaning and disinfection of machinery, equipment and grading lines are, therefore, of primary concern in avoiding spread of the pathogen. However, it is highly likely that other contamination sources may also play an important role in primary infection, e.g. plant debris, alternative hosts (either other crops or weed spp.), irrigation water, aerosols or insects. Potato crops multiplied only twice in the field from pathogen-free mini-tubers in a Dickeya-contaminated area were observed to have c. 30% infection by Dickeya spp. in the harvested tubers (Velvis & van der Wolf, 2008).

Evidence is growing that Dickeya spp. are present in waterways of some countries and that the use of these as a source of irrigation could spread the pathogen to potato fields; thus limiting irrigation from these sources may reduce contamination and disease in potato fields. There is also evidence that alternative hosts could be important in the spread of disease caused by both Dickeya spp. and Pectobacterium spp., so that monitoring/controlling these hosts could help to reduce the incidence of disease (Toth et al., 2006). Besides the use of diagnostics to test stocks of seed tubers for the presence of the pathogen, other control measures effective against the spread of the pathogen within such lots include avoiding poorly drained fields; de-sprouting at planting and over-irrigation; removal of volunteers and potential host plants; removing diseased plants by roguing; rapid haulm destruction; harvesting in dry weather conditions; minimizing damage at harvest and optimizing storage conditions.

Diagnostic methods

Plant protection agencies across Europe are aware of either the presence of Dickeya spp. in their potato production or the potential for its arrival. In many countries, if an attempt is made to differentiate Dickeya spp. and Pectobacterium spp., it is based purely on visual inspection, with few people recording the prevalence of either pathogen (Table 2). As a result, it is usually unclear whether disease is caused by Pectobacterium or Dickeya spp. Diagnostics can effectively be used to identify the presence of Dickeya spp., which can help to avoid planting or exporting Dickeya-infected stocks. Where diagnostics are used, it is mostly intermittent and on a voluntary basis. Although there is not always a clear correlation between Dickeya inoculum levels detected on seed tubers and disease incidence and severity observed after planting (Tsror et al., 2006; Velvis & van der Wolf, 2008), some countries are currently considering using diagnostic testing as a means to differentiate between infections caused by Pectobacterium spp. and Dickeya spp.

A method commonly used for diagnostics is pathogen isolation on selective crystal violet pectate (CVP) medium (Cuppels & Kelman, 1974). In particular, double-layer modifications have recently been used to isolate species of both Pectobacterium and Dickeya (Hyman et al., 2001; Bdliya & Langerfeld, 2005). Incubation on this medium with differential temperatures and erythromycin susceptibility has been proposed for selective isolation of P. atrosepticum, P. carotovorum and Dickeya spp. (Pérombelon & Hyman, 1986) but is not always reliable (Janse & Spit, 1989). A differential medium based on the characteristic production of blue-pigmented indigoidine by Dickeya spp. was recently shown to differentiate Dickeya spp. from soft rot Pectobacterium spp. (Lee & Yu, 2006). On potato dextrose agar (PDA), young colonies of D. dianthicola are either circular, convex, smooth and entire or sculptured with irregular margins, depending on the moisture content of the growth medium. After 4–5 days colonies resemble a fried egg, with a pinkish, round raised centre and lobed periphery, which later becomes feathery or almost coralloid (Lelliott & Stead, 1987). ‘Dickeya solani’ characteristically forms yellow colonies on PDA and some other non-selective media (B. Carter, Fera, York, UK, personal communication).

Serological tests have been used to screen seed potatoes for latent populations of Pectobacterium spp. and Dickeya spp., but have generally been found to lack the required specificity and sensitivity. However, for Dickeya spp. over 10 different serogroups have been identified, and whilst antibodies for P. atrosepticum are generally directed against O-serogroup 1, for Dickeya spp. such antibodies recognize only 68% of isolates (Samson et al., 1990). False positive results and limitations regarding the sensitivity of detection also remain a problem for serological detection of Dickeya spp. (van der Wolf et al., 1993). A monoclonal antibody (6A6) to a fimbrial antigen detected all D. dianthicola isolates tested and some other Dickeya spp. in a triple antibody sandwich (TAS) ELISA (Singh et al., 2000). However, sensitivity was limited to 107 CFU mL−1, compared to a sensitivity of 103 CFU mL−1 for a PCR test using published primers directed to the pectate lyase gene (Nassar et al., 1996). An enrichment ELISA procedure has been used in the Netherlands for routine detection of Dickeya spp. in voluntary testing offered commercially. Advantages include cost efficiency and a correlation of 95% between this method and PCR testing (Nassar et al., 1996; G. W. van den Bovenkamp, NAK, Emmeloord, the Netherlands, personal communication). A new immunoassay based on Luminex xMAP® technology has been proposed as an alternative to ELISA (van der Wolf et al., 2006) for simultaneous detection of P. atrosepticum and Dickeya spp., with pre-enrichment in semiselective polypectate broth helping to achieve the required detection sensitivity.

For routine screening, PCR assays are increasingly used for specific detection and identification of the Dickeya genus (all Dickeya spp. together). The most widely used assays for detection of this group are based on the ADE primers (ADE1/ADE2) from the pectate lyase (pel) gene (Nassar et al., 1996). An alternative PCR method developed by Toth et al. (1999, 2001) allows for the detection of the ‘soft rot erwiniae’ as a single group, together with differentiation of the individual pathogens. Other conventional PCR assays are also available (Smid et al., 1995; van der Wolf et al., 1995). A combined method to detect both the Dickeya group and Patrosepticum was also described based on multiplex PCR, and deriving major advantages in terms of speed (Diallo et al., 2009). The method is reliant on the simultaneous application of Y45/46 primers, described by Fréchon et al. (1998) and which target P. atrosepticum, and primers Ech1/1’ derived from the pelI gene, which target the Dickeya group. Real-time PCR assays based on 16S-23S rDNA sequence were developed in Finland to differentiate Dickeya from Pectobacterium (Laurila et al., 2010) and others are under validation in the UK and the Netherlands. These hope to include D. dianthicola and ‘D. solani’-specific diagnostics, and are based on differences in housekeeping genes or whole genome sequences (J. G. Elphinstone, unpublished data; J. M. van der Wolf, unpublished data). With genome sequencing of representative strains of all Dickeya spp. now underway (L. Pritchard, SCRI, Invergowrie, Dundee, UK, personal communication), it is expected that Dickeya spp.-specific PCR assays will be available in the near future. However, although laborious and expensive, at present the most effective way to identify ‘D. solani’ is to use the method of Nassar et al. (1996) to screen for the Dickeya group, followed by sequencing of either the recA gene (Parkinson et al., 2009) or dnaX gene (Sławiak et al., 2009b). There is also good evidence that REP-PCR and RFLP methods will provide sufficient differentiation to identify these pathogens (Waleron et al., 2002a,b, 2006; Tsror et al., 2009).

The various Dickeya spp. can be routinely identified according to either their fatty acid methyl ester (FAME) profiles (Laurila et al., 2008) or REP-PCR product polymorphisms using enterobacterial repetitive intergenic consensus (ERIC) primers (Sławiak et al., 2009b; D. E. Stead, personal communication). Dickeya dianthicola isolates (including biovars 1, 7 and 9) form a unique profile with either method. Isolates from potato in the Netherlands, originally identified as biovar 5, also group within the typical D. dianthicola profiles. Ribotyping has also been used successfully to type strains within species of Dickeya (Nassar et al., 1994).

Conclusions and perspectives

The recent renaming of the pathogenic species E. chrysanthemi to six different Dickeya spp. has been of considerable value in determining more precisely the causative agents of plant diseases caused by this group of pathogens worldwide. Dickeya dianthicola has been the main Dickeya spp. affecting potato in Europe for over 40 years. However, the last 6 years have seen the emergence of a more aggressive species, with the proposed name ‘D. solani’, which is currently responsible for significant potato crop losses. It is a concern that these pathogens are now able to cause disease in both cool, wet conditions, optimal for disease caused by P. atrosepticum, and warmer, dryer conditions (including those found in Israel and North and South Africa, as well as in continental Europe). This may in the future lead to the increased prevalence of blackleg and related diseases over a wider range of weather conditions favourable to disease development. It is also likely that increased spring and summer temperatures, arising as a consequence of climate change, may exacerbate the problem as Dickeya spp., and especially ‘D. solani’, are more aggressive at these higher temperatures.

In addition to climate change, increased trade is playing a major part in the spread of the disease and, whilst the distribution of seed potato tubers may be the main cause of this spread, other plant hosts (particularly ornamentals) are likely also to play a role. Indeed, there is evidence that D. dianthicola and ‘D. solani’ have spread to potato via this route. If this proves to be the case, given the very wide host range of Dickeya spp., there is a high likelihood that other Dickeya pathogens could also be transferred between plant species (including potato, other crops and wild species), a problem that may intensify with the introduction and spread of new plant species and/or the increased use of irrigation water (that may harbour such pathogens), both as a consequence of climate change. Close consideration thus needs to be given to potential infection pathways, as well as the adaptability of these pathogens to other plant hosts, environments and climatic conditions.

Whatever the source of infection, once in a potato crop the effects of Dickeya spp. can be serious but are not easily differentiated from Pectobacterium spp. based solely on symptomatology. It is, therefore, highly likely that disease outbreaks caused by Dickeya spp. have been underestimated in commercial potato production. Diagnostic tools are thus important for identifying the presence and spread of the pathogens. Diagnostics, based on a variety of molecular methodologies, are now being developed to allow differentiation between Dickeya species, which will be of particular use in future epidemiological investigations. Such diagnostics will be essential if new legislation to prevent the spread of Dickeya spp. into Scotland, Israel and other countries, is to be effective. Other control measures currently recommended are largely in line with those used for Pectobacterium spp., but if this disease is to be controlled, research is needed to identify specific measures that are most effective against the threat from Dickeya spp.