Authors: Don Pitcher, Global Invasive Species Team, The Nature Conservancy
- Xanthium strumarium is an annual that produces a conspicuous prickly 'cocklebur' and ranges from 0.5-6.5 ft. (0.2-2 m) in height. The relatively large, linear to oblong waxy cotyledons helps to distinguish this weed in the early stages of development.
- The first true leaves are opposite, all subsequent leaves are alternate. Leaves are triangular to ovate in outline, have stiff hairs, and are approximately 2-6 in. (5.1-15.2 cm) long. Leaves are irregularly lobed with leaf margins that have relatively inconspicuous teeth.
- Inconspicuous, greenish in color, arising from the area between the leaf petioles and the stems and at the ends of the erect stems.
- An elliptic to egg-shaped two-chambered bur, 0.5-1.5 in. (1.3-3.8 cm) long and covered with hooked prickles. Each bur contains two seeds, one that grows during the first year and one that grows a year later. Two prickles that are longer and wider than the remaining prickles project from the tip of the bur.
- Ecological Threat
- Xanthium strumarium is found throughout the United States and is primarily a weed of agronomic and horticultural crops, nurseries, and occasionally pastures.
Common Name: Rough cockle-bur
Xanthium strumarium is a coarse annual herb. The name Xanthium is derived from the Greek XANTHOS, meaning yellow, from the ancient name of some plant, the fruit of which was used to dye the hair that color (Munz and Keck 1973).
Many specific epithets have been applied to Xanthium strumarium, including: orientale, canadense, chinense, occidentale, macrocarpum, longirostre, pennsylvanicum, and oviforme (Holm et al. 1977). The consensus of taxonomic opinion follows Love and Dansereau's (1959) suggestion that these "species" are actually subspecies or varieties of this highly variable weed. They suggest that X. strumarium consists of seven complexes: strumarium, cavanillesii, oviforme, echinatum, chinense, hybrid, and orientale. There is no evidence of any sterility barriers separating the entities of X. strumarium, but intense inbreeding with occasional outbreeding is responsible for the enormous variation which often results in small, local, but unstable taxa (Love and Dansereau 1959). At least seven varieties or subspecies have been described from California, but today these are generally considered part of the cavanillesii (pennsylvanicum) morphological complex (Love and Dansereau 1959). However, McMillan (1975) considers this a separate species, X. californicum Greene.
Xanthium strumarium is distinguished from spiny clotbur (X. spinosum) by its broader cockleburs, more ovoid leaves on long petioles, and lack of spines.
Xanthium strumarium is a common annual weed spread by water, humans, or other animals. Its origin is still being debated, but cocklebur may be a native California species. It is most abundant on moist open sites but is present on a variety of waste places. Cocklebur is toxic to certain animals. It reproduces from seeds that are viable for up to several years. Biological control measures are currently being investigated and may prove effective in the future. Simple mechanical removal prior to flowering is recommended for control. If pulled following flowering, the plants should be burned. Monitoring should be continued on the sites for several years.
Xanthium strumarium is distributed worldwide (53 degrees north to 33 degrees south latitude) but is most common in the temperate zone (Love and Dansereau 1959). It is a serious weed in Australia, India, South Africa, and the Americas.
There has been considerable controversy regarding the origin of cocklebur. Though first described from Europe, it is probably of American origin (Munz and Keck 1973). Love and Dansereau (1959) suggest that the cocklebur subspecies most abundant in North America (cavanillesii) originated in Central America. The dates of its introduction to California are not known, but it may be pre-Columbian.
Cocklebur is often associated with open, disturbed areas, particularly flood-prone areas with good soil moisture (Martin and Carnahan 1982), but it is found in a wide variety of habitats. It frequents roadsides, railway banks, small streams, and riverbanks, as well as the edges of ponds and freshwater marshes and overgrazed pastures. It does not tolerate shading (Sen 1981).
Cocklebur grows on a wide range of soils (sands to heavy clays) and available moisture. On rich soils with abundant moisture and little competition from other plants, it grows tall and luxuriant, forming pure stands. In dry, poor soils, plants may grow to only a few centimeters high, persist through drought, and set seed. The ability to grow under a variety of conditions results in a continuous seed supply, if plants are not controlled (Holm et al. 1977).
Cocklebur withstands partial submergence for six to eight weeks by forming adventitious roots from the submerged portion of the stem. These roots float in water and often get infested with oxygen-producing green algae (Dedogonium) which solves the problem of aeration (Ambasht 1977).
Weaver and Lechowicz (1983) describe two types of cocklebur populations. Populations located along shores or water courses tend to be small, ephemeral, and homogeneous with seed dispersal by wind and water. Populations in ruderal (weedy) habitats, agricultural fields, or waste areas tend to be large, dense, and heterogeneous with tall, vigorous plants producing an abundance of seed. Seed dispersal here is primarily the result of human activities. Both types of populations, however, occupy unstable habitats and are continually shifting to newly disturbed areas.
Cocklebur is an extremely competitive weed in corn, cotton, and soybeans fields, particularly in the southeastern and midwestern U.S. (Miller 1970, Charudattan and Walker 1982). Though not as abundant in California, it is still a serious problem in agricultural areas (Vargas 1984), as well as in recreation areas and along reservoirs (Wright and Schweers 1984). Some plants appears to have allelopathic properties (Cutler 1983).
The burs cause an allergic reaction in some people (Parsons 1973) and are toxic to domestic animals (and perhaps to some wildlife). Poisoning threats are greatest in areas where other, more palatable plants have already been consumed (Holm et al. 1977). Ingesting an amount of seeds equal to only 0.3 percent of an animal's body weight will cause toxicity. Still, this rarely occurs as the spiny burs are not palatable to animals.
However, the cotyledons are palatable and also have the highest toxicity. Poisoning generally results when these are eaten. This situation occurs most at the edges of ponds, lakes, flood plains, or other bodies of water where shallow flooding followed by recession of the waterline occurs. Under such conditions seeds germinate readily, constantly supplying new generations of potentially poisonous seedlings as the water source dries out. Animals are attracted to such areas because of their need for drinking water. The problem is accentuated because Xanthium seeds do have natural dormancy and germinate over long periods of time. Ingestion of an amount of cotyledons equal to 0.75 to 1.5 percent of the animal's body weight will cause toxicity.
Toxicity decreases rapidly as true leaves are formed. Evidence of poisoning appears in about 12 to 48 hours, the symptoms being nausea, vomiting, lassitude, depression, weakened muscles, and prostration. Severe poisoning may result in convulsions and spasmodic running movements. Ruminants may not vomit. Death may occur within a few hours or days. Fatty substances such as milk, lard, or linseed oil have been recommended as antidotes (Kingsbury 1964).
Kaul (1971) includes the following reasons for cocklebur's ability to inhabit such a range of habitats: an effective dispersal mechanism, wide ecological amplitude, heavy output of seeds and high viability and germination under varied environments, high reproductive capacity, large seed size and weight, rapid seedling growth, and a well-developed root system.
Xanthium strumarium is wind-pollinated, self-compatible, and predominately self-pollinated (Love and Dansereau 1959). The staminate heads of X. strumarium are located above the pistillate heads on the main axis and side shoots, an arrangement favoring inbreeding (Weaver and Lechowicz 1983). Moran and Marshal (1978) found the outcrossing rate in natural populations to be 0 to 12%.
In a Quebec experimental garden, individual plants produced from 611 to 1,488 male inflorescences (Weaver and Lechowicz 1983). The 100-150 male florets in each staminate head begin to shed their pollen from a few days before the stigmata are receptive until all female flowers are ripe. The slightest movement of the plant or a gust of wind causes the pollen to rain down over the exposed stigmata of the female flowers immediately below. The pollen of the plant itself is therefore most likely to ensure the fertiliza- tion of its female flowers, and only an accident, a strong wind, or crowded growth, accomplishes cross-fertilization. In Xanthium, inbreeding is thus the rule and outbreeding only an occasional occurrence (Love and Dansereau 1959).
Cocklebur has been widely used as an experimental plant in studies of photoperiod. Love and Dansereau (1959) list 34 articles on Xanthium photoperiod, and many more have been written since that time (Cleland and Ajami 1974). According to Salisbury (1969), X. strumarium is a short-day plant and usually does not flower when day length exceeds 14 hours. However, there is evidence of differences in light response among the complexes, as some plants flower with day lengths as long as 16 hours.
At high latitudes, day length is greater than 14 hours during summer, and therefore, X. strumarium does not flower until late summer, once day length is short enough to stimulate flowering. Seeds mature late under these conditions, usually in early autumn. These differences are considered to represent genetic adaptations of the reproductive system to environmental variables as a result of natural selection (Ray and Alexander 1966). The cotyledons do not play a role in flower induction (Holm et al. 1977).
SEED PRODUCTION AND DISPERSAL
Open grown X. strumarium plants produce 500 to 5,400 burs per plant. The number of fruits produced is dependent upon the amount of vegetative growth at the time of floral initiation. On crowded plants, production is reduced to 71 to 586 burs per plant (Weaver and Lechowicz 1983). Burs are buoyant and will float for up to 30 days (Kaul 1961), thus being easily dispersed to beaches and pastures subject to flooding. The burs also become entangled in animal hair or human clothing. The burs are a serious problem in sheep production areas where they become entangled in the wool, reducing its value (Wapshere 1974a). X. strumarium burs contain a highly toxic substance, carboxyatractyloside, capable of killing hogs, cattle, goats, horses, sheep, and poultry.
SEED VIABILITY AND GERMINATION
Germination of cocklebur seeds has been extensively researched (Crocker 1906, Davis 1930, Katoh and Esashi 1975, Zimmerman and Weis 1983). More than 80% of cocklebur seeds are viable in most populations (Weaver and Lechowicz 1983). Light is not required for germination, but seedlings seldom emerge from seeds lying on the surface or buried more than 15 cm in the soil (Kaul 1965a, Stoller and Wax 1973).
Seeds of Xanthium strumarium have a high moisture requirement for germination and show little germination in soils at less than 75% of field capacity, but they are able to absorb moisture at high osmotic concentrations (Kaul 1968). Cocklebur seed viability decreases over time, and seeds do not survive more than a few years (Wapshere 1974b). Seedlings are unusually large with foliar-type cotyledons that, through early photosynthetic function, enable the young seedling to become quickly established (Polunin 1966). Seedlings may be identified in the cotyledon stage by the presence (below ground) of the persistent bur, which usually remains attached to the seedling (Kingsbury 1964). The species does not reproduce vegetatively (Weaver and Lechowicz 1983).
Xanthium strumarium plants produce seeds of two types (termed somatic polymorphism). Each bur contains two seeds, with the smaller one often pushed upwards toward the beaked end of the fruit. The lower seed has a shorter dormant period and germinates first. Dormancy in Xanthium involves the presence of a different water-soluble germination inhibitor in each seed type, to which the testa are impermeable. The presence of oxygen causes degradation of these two inhibitors and subsequent rupture of the seed coat, but apparently at very different rates in the two types. Thus at least two batches of seeds are present in each generation to assure germination in the event the immediate environment happens to be unsuitable (Redosevich and Holt 1984).
Xanthium strumarium is considered one of the world's worst weeds (Holm et al. 1977). Cocklebur seeds are easily spread, due to their ability to float and to 'hitchhike' on humans and animals. The plants can quickly become dominant in an area because of their prolific seed production and high germination and survival rates.
Control of cocklebur requires active management once it becomes established in an area.
According to Weaver and Lechowicz (1983), young plants of Xanthium strumarium regenerate readily from the lower nodes if trampled, clipped, or otherwise injured. Fruit on older plants or shoots which have been cut or damaged will continue to ripen provided fertilization has occurred prior to the injury. Burs may persist on dead plants for up to 12 months (Parsons 1973). Since plants can regrow, mowing is not an effective control measure for cocklebur.
Physical removal of the plants by hand pulling or hoeing them is effective if done prior to flowering. If left until after seed development, plants should be carefully removed so as not to dislodge the burs, piled, and burned (Parsons 1973).
Burning is an effective means of destroying cocklebur seeds, but prescribed fire has seldom been used for this purpose.
Much work has been done on the insect pests of Xanthium strumarium (Kelly 1931, Wilson 1960, Hare 1977, 1980, Hare and Futuyma 1978, Foote 1984). Hilgendorf and Goeden (1982, 1983) provide good reviews of sap and foliage feeding (phytophagous) insects associated with X. strumarium. They list 60 different species that attack it in different parts of the world. Although many of these also attack cultivated plants, eight feed only on plants in the Heliantheae Tribe (ragweeds and cocklebur).
The insect fauna of Xanthium species in the central U.S. is richer in species and trophically more specialized than in California (Hilgendorf and Goeden 1983). In California, these insects probably switched from ragweed when cocklebur reached the state. Nine insect species feed on cocklebur as immatures.
Several species of insects have been introduced to Australia to control Xanthium strumarium, but results have generally been disappointing (O'Connor 1952, 1960, Wilson 1960, Wapshere 1974b). The most promising control species there appears to be Nupserha antennata Brun., a beetle native to India and Pakistan (Haseler 1970). Insect species associated with X. strumarium have also been studied in Pakistan (Baloch et al. 1968) and India (Wilson 1960).
Hilgendorf and Goeden (1983) suggest that Oedopa sp. nr. capito (Diptera) is probably the only insect species worthy of study as a potential biocontrol agent for Xanthium strumarium. Oedopa is restricted to the genus Xanthium, feeding on its roots. Baloch and Ghani (1969) suggest that a combination of insect species, with different feeding habits, would improve the chance of suppressing Xanthium populations.
Weaver and Lechowicz (1983) list 14 species of fungi that infect Xanthium in the U.S. and Canada. The rust Puccinia xanthii Schw., which occurs throughout the U.S., southern Canada, parts of Europe, and India, is an obligate parasite on species of Xanthium and Ambrosia (Conners 1967, Hasan 1974, Alcorn 1975, Jadhav and Somani 1978). It attacks all aerial parts of the plant except the flowers. Infected plants mature more rapidly than healthy plants and show decreased transpiration, dry weight, bur production, and percent germination (Hasan 1974, Julien et al. 1979). The spores overwinter on dead plant parts. Fungal and bacterial pathogens have had some success in controlling X. strumarium in India (Deshpande 1982). Kalidas (1981) induced rapid wilt in X. strumarium by using phytopathogenic toxins from seven different fungal and bacterial agents. Plant death was evident within 6 to 8 hours with each toxin. Sharma (1981) also describes a powdery mildew that infects cocklebur in India.
Nematodes reported from X. strumarium are Aphelenchoides ritzema-bosi Schmidt (Weaver and Lechowicz 1983) and meloidogyne hapla (Siddiqui et al. 1973). Cuscuta pentagona (dodder) is a higher plant parasite that has been found on cocklebur (Munz and Keck 1973). Orobanche ramosa L. (broom rape) is another parasitic plant found on a variety of cultivated and weedy plants, including Xanthium (Polunin 1966, Munz and Keck 1973).
CONTROL BY GRAZING
Because of its toxicity and unpalatability, grazing is not a viable control method for Xanthium strumarium.
Cocklebur is susceptible to a wide variety of soil- and foliar- applied herbicides commonly used for the control of broad-leaved weeds (Weaver and Lechowicz 1983), but certain Xanthium complexes are more susceptible than others (Anderson 1982). Dr. Jim McHenry (personal communication 1985), of the University of California, Davis, recommends the following herbicides for cocklebur control in California's preserves:
(1) 2,4-D amine, a phenoxy-type herbicide used for broadleaf weed control, should be applied to plants at the 3- to 5-leaf stage of growth. Application should be at the rate of 1 to 1.5 lbs/100 gallons of water, with one quart of surfactant/100 gallons. (Surfactants lower surface tension of the spray and increase the herbicide's effectiveness.) 2,4-D does not affect grasses.
(2) Dicamba (Banvel) is a broad spectrum herbicide used against perennial broadleaf weeds. It may persist in the soil for up to eight weeks. The suggested mixture is 0.5-0.75 lb/100 gallons water, with one quart of surfactant/100 gallons, and an application rate of 0.5-1.5 pints/acre (not to exceed 2 gal/acre in growing season). Dicamba is more selective than 2,4-D.
(3) Bromoxynil (Buctril, Brominal) is a contact herbicide which affects only the plants or portions of a plant actually contacted by the chemical. Therefore, adequate distribution of the chemical over the foliage is essential. Bromoxynil should not be used on grazed lands but is effective in controlling a wide variety of broadleaf weeds, including Xanthium strumarium. The suggested rate of application is 0.56-1.12 kg/ha (Beste 1983).
(4) Selective weed oils. There are several petroleum oils used for weed control. The herbicidal use of oils depends on their chemical and physical properties. Most contact oils evaporate slowly and owe their plant toxicity to their high content of aromatic compounds. Spraying oil on cocklebur will be effective only if entire plants are coated.
Herbicides can be applied uniformly over an area (for large infestations) or by spot spraying individual plants. Dr. McHenry recommends using a flat-fan nozzle (Spraying Systems Co. #8003 or #8004 nozzle tip) rather than the cone nozzles available on most garden sprayers, as cone sprayers produce greater atomization of the chemicals and increase the chance of drift into unwanted areas. Spraying should be done on calm days when plant surfaces are dry.
Management Research Needs
The origin of Xanthium strumarium needs to be determined. Additional research is needed on the possible toxic effects of X. strumarium on wildlife, on biological control measures, and on the effects of prescribed fires.
Abrams, L. 1940. Illustrated flora of the Pacific states: Washington, Oregon, and California. Vol. I. Ophioglossaceae to Aristolochiaceae. Stanford Univ. Press, Stanford, California. 538 pp.
Alcorn, J. L. 1975. A new disease of Noogoora burr. Queensland Agricultural J. 101:162.
Ambasht, R. S. 1977. Observations on the ecology of noxious weeds on Ganga River banks at Varanasi, India. Vol. 1, p. 109-115 in 6th Asian-Pacific Weed Science Society Conference, Indonesia. 365 pp.
Anderson, R. N. 1982. Variation in growth habit and response to chemicals among three common cocklebur (Xanthium strumarium) selections. Weed Science 30:339-343.
Baloch, G. M. and A, I. Mohyuddin, and M. A. Ghani. 1968. Xanthium strumarium L.--insects and other organisms with it in West Pakistan. Commonwealth Institute Biological Control Technical Bulletin 10:103-111.
Baloch, G. M. and M. A. Ghani. 1969. The present status of biological control of Xanthium (Compositae). PANS 15:154-159.
Beste, C. E. 1983. Herbicide handbook. Weed Science Society of America, Herbicide Handbook Committee. Champagne, IL.
Charudattan, R. and H. L. Walker. 1982. Biological control of weeds with plant pathogens. John Wiley & Sons, New York. 293 pp.
Cleland, C. F. and A. Ajami. 1974. Identification of the flower-inducing factor isolated from aphid honeydew as being salicylic acid. Plant Physiology 54:904-906.
Cole, R. J., B. P. Stuart, J. A. Lansden, R. H. Cox. 1980. Isolation and redefinition of the toxic agent from cocklebur (Xanthium struamrium). J. Agric. Food Chemistry 28:1330-1332.
Conners, I. L. 1967. An annotated index of plant diseases in Canada and fungi recorded on plants in Alaska, Canada and Greenland. Can Dept. Agric. Publ. 1251. Ottawa, Ont. 381 pp.
Crocker, W. 1906. Pole of seed coat in delayed germination. Botanical Gazette 42:265-291.
Crockett, L. J. 1977. Wildly successful plants. McMillan Publishing Co., New York. 268 pp.
Cutler, H. G. 1983. Carboxyatractyloside: a compound from Xanthium strumarium and Atractylis gummifera with plant growth inhibiting properties. The probable "Inhibitor A." J. Natural Products 46:609-613.
Davis, W. E. 1930. The development of dormancy in seed of cocklebur (Xanthium). American Journal of Botany 17:77-87.
Deshpande, K. 1982. Biocontrol of Parthenium hysterophorous L. and Xanthium strumarium L. through phytopathogens. In: Abstracts of papers, annual conference of Indian Society of Weed Science. p. 48.
Foote, B. A. 1984. Host plant records for North American ragweed flies (Diptera: Tephritidae). Entomological News 95:51-54.
Hare, D. J. 1977. The biology of Phaneta imbridana (lepidoptera: Tortricidae) a seed predator of Xanthium strumarium (Compositae). Psyche 84:179-182.
Hare, D. J. 1980. Variation in fruit size and susceptibility to seed predation among and within populations of the cocklebur, Xanthium strumarium L. Oecologia 46:217-222.
Hare, D. J. and D. J. Futuyma. 1978. Different effects of variation in Xanthum strumarium L. (Compositae) on two insect seed predators. Oecologia 37:109-120.
Hasan, S. 1974. Recent advances in the use of plant pathogens as biocontrol agent of weeds. Pest Articles and New Summaries 20:437-443.
Haseler, W. H. 1970. Insects in Noogoora burr control. Queensland Agriculture Journal 96:191-193.
Hilgendorf, H. H. and R. D. Goeden. 1983. Phytophagous insect faunas of spiny clotbur, Xanthium spinosum, and cocklebur, Xanthium strumarium. Environmental Entomolgy 12:404-411.
Hilgendorf, J. H. and R. D. Goeden. 1982. Phytophagous insects reported worldwide from the noxious weeds spiny clotbur, Xanthium spinosum, and cocklebur, Xanthium strumarium. Entomological Society of America 28:147-152.
Holm, L. G., P. Donald, J. V. Pancho, and J. P. Herberger. 1977. The World's Worst Weeds: Distribution and Biology. The University Press of Hawaii, Honolulu, Hawaii. 609 pp.
Jadhav, A. N. and R. B. Somani. 1978. Puccinia Xanthii--a report from India. Indian Phytopathology 31:369-371.
Jepson, W. L. 1951. Manual of the flowering plants of California. University of California Press, Berkeley.
Julien, M. H., J. E. Boradbent, and N. C. Matthews. 1979. Effects of Puccinia xanthii on Xanthium strumarium (Compositae). Entomophaga 24:29-34.
Kalidas, D. 1981. Phytopathogens as weed control agents. Proceedings 8th Asian-Pacific Weed Science Society Conference, pp. 157-159.
Katoh, H. and Y. Esashi. 1975. Dormancy and impotency of cocklebur seeds. I. CO2, C2H4, O2, and high temperature. Plant and Cell Physiology 16:72-87.
Kaul, V. 1961. Water relations of Xanthium strumarium L. Sci. Cult. 27:495-497.
Kaul, V. 1965. Physiological-ecology of Xanthium strumarium L. I. Seasonal morphological variants and distribution. Tropical Ecology 6:72-87.
Kaul, V. 1965. Physiological-ecology of Xanthium strumarium L. II. Physiology of seeds in relation to its distribution. J. Indian Bot. Soc. 44:365-380.
Kaul, V. 1968. Physiological-ecology of Xanthium strumarium L. V. Water relations. Tropical Ecology 9:88-102.
Kaul, V. 1971. Physiological-ecology of Xanthium strumarium L. IV. Effect of climatic factors on growth and distribution. New Phytologist 70:799-812.
Kelly, S. G. 1931. The control of Noogoora and Bathurst burr by insects. J. Counc. Sci. Ind. Res. (Australia) 4:161-172.
Kingsbury, J. M. 1964. Poisonous plants of the U.S. and Canada. Prentice-Hall, Inc., Englewood Cliffs, NJ. 626 pp.
Love, D. and P. Dansereau. 1959. Biosystematic studies on Xanthium: toxonmic appraisal and ecological status. Candian J. Botany 37:173-208.
Martin, R. J. and J. A. Carnahan. 1982. Distribution and importance of Noogoora and Bathurst burrs in eastern Australia. Austrailian Weeds 2:27-32.
McHenry, Jim. 1985. Extension Weed Scientist, University of California, Davis, Cooperative Extension, CA. Personal communication. May 1985.
McMillan, C. 1975. The Xanthium strumarium complexes in Australia. Australian J. Botany 23:173-192.
Miller, J. F. 1970. Cocklebur. Crops and Soils 22:15-17.
Moran, G. F. and D. R. Marshal. 1978. Allozyme uniformity within and variation between races of the colonizing species Xanthium strumarium L. (Noogoora burr). Australian J. Biological Science 31:283-291.
Munz, P.A., and D.D. Keck. 1973. A California flora and supplement. Univ. California Press, Berkeley, CA.
O'Connor, B. A. 1952. An introduced parasite of Noogoora burr. Agricultural Journal, Department of Agriculture, Fiji 23:105-106.
O'Connor, B. A. 1960. A decade of biological control work in Fiji. Agricultural Journal, Department of Agriculture, Fiji 30:44-54.
Parsons, W. T. 1973. Noxious weeds of Victoria. Inkata Press, Ltd., Melbourne, Australia. 300 pp.
Polunin, N. (ed.) 1966. Weeds of the world: biology and control. Gramian Press, Inc., London. 526 pp.
Ray, P. M. and W. E. Alexander. 1966. Photoperiodic adaption to latitude in Xanthium stumarium. American Journal of Botany 53:806-816.
Redosevich, S. R. and J. S. Holt. 1984. Weed ecology. John Wiley & Sons, New York.
Robbins, W. W. 1940. Alien plants growing without cultivation in California. California Agricultural Experiment Station Bulletin 637:1-128.
Robbins, W.W., M.K. Bellue, and W.S. Ball. 1970. Weeds of California. State of California, Department of Agriculture. 547 pp.
Salisbury, F. B. 1969. Xanthium strumarium L. pp. 14-16 in: L. Evans, ed. The induction of flowering: some case histories. MacMillan, Melbourne. 488 pp.
Sen, D. N. 1981. Ecological approaches to Indian weeds. Geobios International, Jodhpur, India. 301 pp.
Sharma, A. K. 1981. The powdery mildew of Xanthium strumarium from J and K state. Indian Journal of Mycology and Plant Pathology 11:92-95.
Siddiqui, I. A., A. A. Sher, and A. M. French. 1973. Distribution of plant parasitic nematodes in California. California Department of Food and Agriculture, Sacramento. 324 pp.
Stoller, W. W. and L. M. Wax. 1973. Periodicity of germination and emergence of some annual weeds. Weed Science 21:574-580.
Vargas, R. 1984. Weed management systems for cotton. Pp.52-56 in: Proceedings 36th Annual California Weed Conference. 164 pp.
Wapshere, A. J. 1974. An ecological study of an attempt at biological control of Noooora burr (Xanthium strumarium). Australian J. Agricultural Research 25:275-92.
Wapshere, A. J. 1974. The regions of infestation of wool by Noogoora bur (Xanthium strumarium), their cliamtes and the biological control of the weed. Australian J. Agricultural Research 25:775-81.
Weaver, S. E. and M. J. Lechowicz. 1983. The biology of Canadian weeds. 56. Xanthium strumarium L. Canadian J. Plant Science 63:211-225.
Wilson, F. 1960. A review of the biological control of insects and weeds in Australia and Australian New Guinea. Commonwealth Institute Biological Control Technical Communication No. 1:1-102.
Wright, S. D. and V. H. Schweers. 1984. Control of cocklebur at Lake Success, California. Proceedings of the Western Society of Weed Science 37:215-217.
Zimmerman, J. K. and I. M. Weis. 1983. Fruit size variation and its effects on germination and seedling growth in Xanthium strumarium. Canadian J. Botany 61:2309-2315.