Orobanche minor
(common broomrape)

Pictures

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Identity

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Preferred Scientific Name

• Orobanche minor Sm. (1797)

Preferred Common Name

• common broomrape

Other Scientific Names

• Orobanche abyssinnica A. Rich. (1851)
• Orobanche apiculata Wallr. (1822)
• Orobanche barbata Poir.
• Orobanche concolor Duby
• Orobanche crithmi Bertol.
• Orobanche euglossa Rchb.f.
• Orobanche grisebachii Reut. 1846
• Orobanche hyalina Spruner ex Reut.
• Orobanche laurina Bertol.
• Orobanche livida Sendtn. ex Freyn
• Orobanche pyrrha Rchb.f.
• Orobanche unicolor Boreau
• Orobanche yuccae Pa.Savi ex Bertol.

International Common Names

• English: clover broomrape; lesser broomrape; small broomrape
• Spanish: rabo de lobo
• French: Orobanche du trefle; petit orobanche
• Portuguese: erva-toira-menor

Local Common Names

• Germany: Kleeteufel; Kleine sommerwurz
• Italy: Orobanche del trifoglio
• Japan: yase-utsubo
• Netherlands: klavervreter
• USA: hellroot

EPPO code

• ORAGB (Orobanche grisebachii)
• ORAMI (Orobanche minor)

Summary of Invasiveness

Top of page Orobanche minor is not a highly invasive plant but its introduction could result in a significant impact on certain crop species, especially forage legumes and tobacco.

Taxonomic Tree

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• Domain: Eukaryota
•     Kingdom: Plantae
•         Phylum: Spermatophyta
•             Subphylum: Angiospermae
•                 Class: Dicotyledonae
•                     Order: Scrophulariales
•                         Family: Orobanchaceae
•                             Genus: Orobanche
•                                 Species: Orobanche minor

Notes on Taxonomy and Nomenclature

Top of page O. minor is in the section Osproleon of the genus Orobanche (together with species including O. crenata and O. cernua) with simple bracts, whereas the other important section Trionychon (containing O. ramosa and O. aegyptiaca) has additional bracteoles on the calyx.

The taxonomy and nomenclature of O. minor is a matter of some disagreement as it exists in a wide range of forms which are variously treated as varieties, sub-species or distinct species. Beck-Mannagetta (1930) recognises two varieties, var. genuine and var. neglecta, and many 'forma' but Flora Europaea (Chater and Webb, 1972) recognises no divisions. Benharrat et al. (2000) have recently used molecular techniques to confirm the distinctions between O. minor and the closely related taxa O. amethystea, O. loricata and O. hederae. Stace (1991) and Rumsey and Jury (1991) list varieties in the UK as: var. minor (on a wide range of hosts); var. flava with yellow corolla (on Hypochaeris radicata and related species); var. maritima with yellow bosses on the lower lip (on Daucus carota, Plantago coronopus and Ononis repens); and var. compositarum, with narrow erect corollas pressed to the stem (mainly on Crepis virens [C. capillaris] and other Compositae [Asteraceae]). Where morphological variants are associated with particular host species it is not known whether the morphology is affected by the host, nor whether these variants are truly host-specific.

Description

Top of page Orobanche species are obligate parasites on the roots of various host plants and lack any chlorophyll. After germination and attachment to a host root, a tuberous 'nodule' develops, usually covered in 'crown roots', short roots up to 1-2 cm long of uncertain function. Some weeks after germination and development, the shoot erupts from this tuberous mass. In O. minor the stems are un-branched, yellowish-brown, often tinged with purple, generally 30-50 cm tall but sometimes exceeding 100 cm (especially in Ethiopia), glandular-villous or nearly glabrous. Leaves are represented by alternate brown scales, ovate to lanceolate, acuminate, 6-20 mm long. The inflorescence is an elongated spike, 10-30 cm long (longer in very large specimens) occupying about half the mature stem. Flowers are sessile, arranged spirally, each subtended by a single narrow, ovate-acuminate, glandular-hairy bract 10-15 mm long x 3-5 mm wide. The calyx, also glandular-hairy, is variable in form, usually with two pairs of acute, almost subulate lobes divided to about halfway laterally, but sometimes with only two undivided lobes. The lobes, or pairs of lobes, are completely separated by deep sinuses dorsally and ventrally. The corolla is 10-18 mm long, rarely to 20 mm, the tube more-or-less cylindrical, slightly curved downwards, opening out to lobes 2-3 mm long, to make a total width of about 10 mm at the mouth, glabrous to moderately glandular-hairy on the outside, generally glabrous within. Colour mainly pale whitish but with varying amounts of purple concentrated along the veins. Stamens inserted 2-4 mm from the base of the corolla tube, the filaments often hairy towards the base. Stigma 2-lobed, reddish-brown, rarely yellow. Capsule 7-10 mm long, splitting into two valves when ripe, shedding several hundred minute seeds, about 0.3 mm long with coarse reticulate marking.

Distribution

Top of page O. minor has a wide natural distribution through most of Europe (excluding the extreme north), North Africa and western Asia. It has been introduced further afield in the USA, S. America, Australia, New Zealand and southern Africa. It is less clear whether occurrence in the highland tropics of eastern Africa is natural or introduced.

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

History of Introduction and Spread

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Introduced to the USA, Japan, Chile, Australia, New Zealand, and southern Africa but little available information on the dates or methods of introduction. It has not proved invasive in the USA but causes concern in South Africa (Henderson et al., 1987) and has been a serious pest of tobacco locally in New Zealand (James and Frater, 1977).

Frost and Musselman (1980) report that in a survey of herbarium material, O. minor was found to have a 116-year history in the USA. The species has been collected 98 times in 12 states plus the District of Columbia and appears to be established at maritime ports and along railway lines. The distribution pattern suggests repeated introductions from foreign sources and persistence in some areas. Most of the infestations in the USA have later been eradicated, either by removal of host plants or by fumigation. (Eplee et al., 1994).

In Chile, O. minor is known to have occurred since 1952 and has spread south to latitude 39°S from its original site at latitude 37°S (Norambuena et al., 1999).

In Japan, O. minor was first recorded in 1937. It has spread to much of the country but has not apparently caused serious concern (K. Yoneyama, Utsunomiya University, Utsunomiya 321-8505, Japan, personal communication).

Risk of Introduction

Top of page The risk of introduction is not high, as O. minor is not an especially frequent weed of crops and, even when it is, the chances of contamination of crop seed is moderate or low. However, any contamination that does occur may have serious consequences where it contravenes plant quarantine regulations.

Habitat

Top of page O. minor is associated mainly with disturbed areas, crops, roadsides and waste places on a wide range of soil types in temperate and highland tropical areas. There is some tendency to association with alkaline soils but the local importance of O. minor on tobacco suggests that this is not a strict limitation. Although the other main group of root parasites, Striga spp., are favoured by low soil nitrogen, this is not generally true for Orobanche spp. Yoneyama et al. (2001) even report increased stimulation of O. minor germination by exudates from roots of Trifolium pratense at increasing levels of nitrate, although ammonium could be inhibitory. Phosphate also greatly reduced stimulant exudation, in keeping with field observations of reduced O. minor infestation in Trifolium subterraneum at higher phosphate levels (Southwood, 1971). Westwood and Foy (1999) also showed ammonium nitrogen to be directly inhibitory to radicle elongation in a number of Orobanche species, O. minor being the most susceptible to this effect. O. minor may occur in some irrigated crops but is not common in naturally wet habitats.

Habitat List

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Category	Sub-Category	Habitat	Presence	Status
Terrestrial
	Terrestrial – Managed	Cultivated / agricultural land	Present, no further details	Harmful (pest or invasive)
		Managed grasslands (grazing systems)	Present, no further details	Harmful (pest or invasive)
		Disturbed areas	Present, no further details	Harmful (pest or invasive)
		Rail / roadsides	Present, no further details	Harmful (pest or invasive)

Hosts/Species Affected

Top of page O. minor parasitizes numerous species in a wide range of plant families, predominantly Leguminosae [Fabaceae], Umbelliferae, Solanaceae and Compositae [Asteraceae] but also many others including Malvaceae, Geraniaceae and Rosaceae. Musselman et al. (1981) found some evidence for the existence of biotypes with differing specificity; a population from a lettuce host parasitized tobacco and Trifolium species as well as lettuce, whereas populations from Trifolium species failed on lettuce and were very slow to emerge on tobacco. However, seeds of O. minor var. minor taken from plants parasitizing Eryngium maritimum were able to parasitize Hypochaeris radicata, Trifolium hybridum and T. pratense (Hipkin, 1992).

Where morphological variants are associated with particular host species it is not known whether the morphology is affected by the host, nor whether these variants are truly host-specific.

Growth Stages

Top of page Flowering stage, Fruiting stage, Vegetative growing stage

Latitude/Altitude Ranges

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Air Temperature

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Rainfall

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Natural enemies

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Notes on Natural Enemies

Top of page By far the most significant insect enemy of O. minor is the dipteran Phytomyza orobanchia, associated with O. minor through most if not all of its natural range. Only a selection of countries are indicated in the Table. Kroschel and Klein (1999) indicate its occurrence in most of Europe, the Mediterranean, the Balkans and Central Asia. Reductions of total seed production resulting from natural infestations range from 11 to 90% and it has been exploited for biological control (see 'Control' section).

Orobanche species are affected by a wide range of fungal pathogens and many of those recorded by Linke et al. (1992) on O. crenata are likely to also occur on O. minor, in addition to Aspergillus niger and Alternaria alternata which were recorded on O. minor. The complete collapse of plants sometimes observed after attack by Phytomyza is often the result of fungal infection associated with the physical damage caused to the stems by Phytomyza larvae.

Means of Movement and Dispersal

Top of page Natural Dispersal (Non-Biotic)

Seeds of O. minor are small enough to be blown some distance by wind but rarely further than a few metres. Movement in surface water is likely to occasionally provide longer-distance dispersal.

Vector Transmission (Biotic)

Movement by animals may occasionally occur but only in a sporadic way.

Agricultural Practices

By far the most important agent of local dispersal is farm machinery transporting contaminated soil from field to field and from farm to farm.

Accidental Introduction

Accidental introduction over long distances, including national boundaries, may occur when contaminated soil or crop seed is introduced.

Intentional Introduction

Intentional introduction is unlikely.

Plant Trade

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Plant parts not known to carry the pest in trade/transport
Bark
Seedlings/Micropropagated plants
Wood

Impact Summary

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Impact

Top of page The effects of Orobanche species on their hosts are somewhat less severe than those of Striga species. Damage to the host in the studies reported so far generally appears to be proportional to the dry weight of the parasite, so that the total dry weight of the host/parasite combination equals the weight of the un-infested host. Damage from O. crenata on faba bean may be greater than this under dry conditions, apparently due to some enhancement of drought stress, but this has not been reported for O. minor. Apart from yield reduction, O. minor can greatly reduce the value of clover and lucerne seed crops due to the danger of seed contamination. In Chile, it is seriously harming exports of Trifolium pratense seed and 6,000 ha on 500 farms are potentially endangered (Norambuena at al., 1999). It has been reported as killing ornamental species including Pelargonium x hortorum and Campanula portenschlagiana in Ireland (Murphy, 1972).

O. minor is recorded by Holm et al. (1979, 1997) as a 'serious' weed of beans and peas in Egypt, of legumes in Czechoslovakia, and of clover, rape and tobacco in New Zealand; also a 'principal' weed of legumes in Italy and Pakistan, and in grassland in Uganda. It is also common in lucerne in Turkey, in legumes in Hungary and vegetables in Saudi Arabia. The most widespread hosts appear to be the forage legumes (Trifolium spp., Medicago sativa and Lotus corniculatus). In Ethiopia it may cause local damage to safflower, sunflower, tobacco, noug (Guizotia abyssinica), lettuce, groundnut, faba bean and tobacco, as well as to clovers (Parker, 1992).

In some countries, the importance of O. minor as a weed has declined with the reduced cultivation of Trifolium seed crops, as in the Netherlands (ter Borg et al., 1994), and in the UK (Parker and Riches, 1993).

Environmental Impact

Top of page No significant environmental impact has been reported for O. minor.

Impact: Biodiversity

Top of page No significant impact on biodiversity has been reported for O. minor.

Social Impact

Top of page No significant social impact has been reported for O. minor.

Risk and Impact Factors

Top of page Invasiveness

• Invasive in its native range
• Proved invasive outside its native range
• Highly mobile locally
• Has high reproductive potential
• Has propagules that can remain viable for more than one year

Impact outcomes

• Negatively impacts agriculture

Impact mechanisms

• Competition - monopolizing resources

Likelihood of entry/control

• Highly likely to be transported internationally accidentally
• Difficult to identify/detect as a commodity contaminant
• Difficult/costly to control

Uses

Top of page No uses of this species are known.

Similarities to Other Species/Conditions

Top of page O. minor is distinguished from the important weedy species O. ramosa and O. aegyptiaca by lack of branching and the absence of bracteoles. There is greater difficulty distinguishing it from other members of the section Osproleon, especially from dried material. O. crenata is close to O. minor in many respects but is larger in most of its parts and its flowers are almost invariably over 20 mm long and over 12 mm across. In O. cernua (including O. cumana), the flowers are similar in size to those of O. minor, but when fresh they are clearly distinguished by the contrast between a white tube and deeply coloured lobes. Dried material shows less contrast, but dissection of the corolla tube shows the stamens of those two species to have glabrous filaments inserted at least 5 mm from the base of the tube.

Benharrat et al. (2000) have recently used molecular techniques to confirm the morphological distinctions between O. minor and the closely related taxa O. amethystea, O. loricata and O. hederae. For a full monograph see Beck-Mannagetta (1930). For keys which include all the main species of concern see Chater and Webb (1972) and Parker and Riches (1993).

Prevention and Control

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Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Cultural Control

Infestation may be prevented by phytosanitation, aimed at preventing the spread of viable seeds. This may involve minimizing the movement of infested soil by farm machinery and vehicles, preventing grazing on infested plant material, treating manure (e.g. composting) and avoiding the use of hay made from Orobanche-infested plants (Jacobsohn, 1984). The use of Orobanche-infested crop seeds should also be avoided.

Trap crops and catch crops are used in the control of some other Orobanche species. These stimulate germination of Orobanche in the soil, in order to deplete the seed reserve. Trap crops promote Orobanche seed germination but do not support parasitism; catch crops support parasitism but are destroyed prior to Orobanche flowering. Examples of trap crops for Orobanche include flax (Ramaiah, 1987), mung beans, maize and sorghum (Foy et al., 1989). A catch crop that has been effective in Egypt is berseem (Trifolium alexandrinum), harvested repeatedly for forage to prevent full development and seeding of the parasite (Al-Menoufi, 1994). Kropac (1973) has also reported that a catch crop of Trifolium repens and Lolium multiflorum was effective in eliminating seeds of Orobanche spp., including O. minor. Though these methods have in some cases proved successful, the use of trap and catch crops has limitations for two reasons: (a) local strains of the parasites may differ in their response; (b) it takes several years of trap or catch cropping to reduce parasite seed populations to non-damaging levels.

Frater (1975) proposes deep ploughing as a control measure, depending on the observation that infestation on tobacco in New Zealand arises from shallow soil layers.

Fertilization, especially phosphate and ammonium nitrogen, can help to suppress O. minor (Yoneyama et al., 2001). Westwood and Foy (1999) show that the levels of ammonium required to inhibit O. minor are non-toxic to a range of likely host crops.

Host-plant resistance is an important approach to the control of several other species of Orobanche, but there is no known instance of the selection of crop varieties resistant to O. minor.

Mechanical Control

Hand-pulling of Orobanche flowering stems can be useful, preferably at an early stage before maximum damage has been caused, but stems should immediately be removed from the field as they may continue developing flowers and spreading seeds even without being connected to the host. Hand-weeding is very important, especially when only a few Orobanche plants develop in a field. This can prevent further spread of the parasite and avoid damage.

Chemical Control

Soil fumigation with metham sodium [metam] has been used with moderate success against O. minor in New Zealand (James and Frater, 1977).

There are few herbicides that have an adequate margin of selectivity against Orobanche species and none have been developed specifically for control of O. minor. Glyphosate can be used for control of O. crenata in faba bean and some other crops if applied at low rates (Garcia-Torres, 1994). The imidazolinone herbicides have shown promise applied in various ways for control of Orobanche spp. in legumes and sunflower, especially imazethapyr pre-emergence, post-emergence or as seed dressings in legumes, and imazapyr in lentil and sunflower (e.g. Garcia-Torres and Lopez-Granados, 1991a, b; Garcia Torres et al., 1995, 1996, 1998). A number of these options are likely to be suitable for control of O. minor in the relevant crops, but these compounds will need to be used with care to avoid the development of herbicide-resistant populations of Orobanche. For a broad review of control options for Orobanche spp., see Parker and Riches (1993).

The use of transgenic crops engineered with target-site herbicide resistance is perhaps one of the most promising solutions for Orobanche infestation in many crops. Using glyphosate on transgenic oilseed rape, and chlorsulfuron and asulam on tobacco, complete control of O. aegyptiaca was achieved without affecting the crop or its yield (Joel et al., 1995). These techniques have yet to be applied to the control of O. minor, but when the relevant herbicide-resistant crops become available they should prove valuable.

Biological Control

The broomrape-fly Phytomyza orobanchia was widely used for Orobanche control in the Soviet Union and some East European countries in the 1960s and 1970s (Girling et al., 1979). Infested Orobanche shoots were collected during the growing season, dried and stored under controlled temperature and humidity over the winter and released from specially constructed 'phytomyzaria' which allowed separation and destruction of the smaller hyperparasites as they emerged. The topic has recently been reviewed in some detail and there has been some renewed interest in its exploitation (see Kroschel and Klein, 1999; Klein and Kroschel, 2002). These authors provide detailed reviews of the biology of the insect, its very large number of hyper-parasites (at least 24 are listed) and the early elaborate systems for its exploitation in eastern Europe, where success was often over 90% in terms of reduced Orobanche seed production in the treated fields, but problems had arisen from interference by hyperparasites and also from insecticide use. They emphasise that repeated releases are needed over several seasons to overcome the problem of the long-lived seed bank. Recent attempts at introduction of P. orobanchia to Chile as a classical biocontrol agent against introduced infestations of O. minor and O. ramosa are showing promise (Normabuena et al., 1999, 2001).

Orobanche-specific forms of Fusarium oxysporum have shown promise for control of some other Orobanche spp. in tobacco and in sunflower (Parker and Riches, 1993). Ulocladium atrum has also appeared promising against O. crenata (Linke et al., 1992), but neither of these agents has yet been developed for practical use.

Integrated Control

Ideas for integrated control of Orobanche species are reviewed by Pieterse et al. (1992) and by Parker and Riches (1993). Approaches for possible inclusion in an integrated control programme include general techniques of crop rotation, flooding, fumigation, solarization, prevention of seeding and biocontrol, as well as more crop-specific methods including crop resistance, herbicides and time of planting.

References

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Benharrat H, Delavault P, Theodet C, Figureau C, Thalouarn P, 2000. rbcL Plastid pseudogene as a tool for Orobanche (subsection Minores) identification. Plant Biology, 2(1):34-39; 34 ref.

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Dorr I, Kollman R, 1975. Structural features of parasitism of Orobanche. 2. Differentiation of assimilate conducting elements within the haustorium. Protoplasma, 83(3):185-201

Eplee RE, Fay CL, Norris R, 1994. Orobanche infestations in the USA. Biology and management of Orobanche. In: Pieterse AH, Verkleij JAC, Borg SJ ter, eds. Proceedings of the third international workshop on Orobanche and related Striga research, Amsterdam, Netherlands, 8-12 November 1993. Amsterdam, Netherlands: Royal Tropical Institute, 611-613

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Frost CC, Musselman LJ, 1979. Studies of the distribution and floral biology of Orobanche minor. Proceedings of the 2nd International Symposium on Parasitic Weeds, North Carolina, 1979., Supplement:3

Frost CC, Musselman LJ, 1980. Clover broomrape (Orobanche minor) in the United States. Weed Science, 28(1):119-122

Garcia Torres L, 1994. Progress in Orobanche control, an overview. In: Pieterse AH, Verkleij JAC, Borg SJ ter, eds. Biology and management of Orobanche. Proceedings of the 3rd International Workshop on Orobanche and Related Striga Research, Amsterdam, Netherlands: KIT, 390-399.

Garcia-Torres L, Castejon-Munoz M, Lopez-Granados F, Jurado-Exposito M, 1995. Imazapyr applied postemergence in sunflower (Helianthus annuus) for broomrape (Orobanche cernua) control. Weed Technology, 9:819-824.

Garcia-Torres L, Jurado-Exposito M, Castejon-Munoz M, Lopez-Granados F, 1996. Herbicide-treated crop seeds for control of Orobanche spp. In: Moreno MT, Cubero JI, eds. Advances in Parasitic Plant Research, Sevilla, Spain: Junta de Andalucia, 699-715.

Garcia-Torres L, Lopez-Granados F, 1991. Control of broomrape (Orobanche crenata Forsk.) in broad bean (Vicia faba L.) with imidazolinones and other herbicides. Weed Research (Oxford), 31(4):227-235

Garcia-Torres L, Lopez-Granados F, 1991. Progress of herbicide control of broomrape (Orobanche spp.) in legumes and sunflower (Helinanthus annuus L.). In: Ransom JK, Musselman LJ, Worsham AD, Parker C, eds. Proceedings of the Fifth international symposium of parasitic weeds, Nairobi, Kenya, 24-30 June 1991. Nairobi, Kenya: CIMMYT (International Maize and Wheat Improvement Center), 306-309

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Jones M, 1991. Studies on the pollination of Orobanche species in the British Isles. Progress in Orobanche research. Proceedings of the international workshop on Orobanche research, Obermarchtal, Germany, 19-22 August 1989., 6-17; 18 ref.

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Kropac Z, 1973. Weedy Orobanche spp. of Czechoslovakia and the range of their parasitism. Symposium on Parasitic Weeds, Malta, 35-43.

Kroschel J, Klein O, 1999. Biological control of Orobanche spp. with Phytomyza orobanchia Kalt., a review. In: Kroschel J, Abderabihi M, Betz H. eds. Advances in Parasitic Weed Control at On-farm Level, Volume II. Wekersheim, Germany: Margraf Verlag, 135-159.

Linke KH, Schneibel C, Saxena MC, Sauerborn J, 1992. Fungi occurring on Orobanche spp. and their preliminary evaluation for Orobanche control. Tropical Pest Management, 38:127-130.

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Musselman LJ, Parker C, 1982. Preliminary host ranges of some strains of economically important broomrapes (Orobanche). Economic Botany, 36(3):270-273

Musselman LJ, Parker C, Dixon N, 1981. Notes on autogamy and flower structure in agronomically important species of Striga (Scrophulariaceae) and Orobanche (Orobanchaceae). Beitrage zur Biologie der Pflanzen, 56:329-343.

Norambuena H, Diaz J, Kroschel J, Klein O, Esacobar S, 2001. Rearing and field release of Phytomyza orobanchia on Orobanche ramosa in Chile. In: Fer A, Thalouarn P, Joel DM, Musselman LJ, Parker C, Kerkleij, eds. Proceedings of the Seventh International Parasitic Weed Symposium, Nantes, 2001, 258-261.

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