ID large-thorned plant growing in Turkey

ID large-thorned plant growing in Turkey

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Can anyone identify this plant which is growing in a friend's garden in the Uzumlu area of Turkey. It has very large spikes/thorns. I'm told it doesn't have flowers or seeds so we are wondering how it propagates itself.

This is a palm (family Arecaceae). Many palms have spines; I'm no expert but it looks to me like it is a member of the Phoenix genus, perhaps a Cretan date palm (Phoenix theophrasti). Date palms do propagate themselves by seed (the pit of the date fruit). The palm in your picture, if a date palm, would be rather young, which could explain why it does not presently show flowers or fruits. Here is a photo of the spines and flowers of a Cretan date palm (by Wouter Hagens, from Wikimedia Commons):

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Food Plot Fertilizers and Management

Put the Taxidermist on Speed Dial

BioLogic has changed the way many Gamekeepers manage their properties. Our blends like Maximum which are 100% NZ brassicas are capable of growing TEN tons of forage per acre, as opposed to traditional cereal grains that might grow a single ton. BioLogic blends can feed a lot of deer. That's our magic. We overwhelm your deer herd with nutritious forages, and soon your neighbors deer show up. However, you have to do all this right. 1. Plant fall plots as early as possible (late summer if weather permits) 2. Plant at the correct depth and at the recommended rate (more is not better) 3. Fertilize accordingly 4. Soil test prior to next season and add missing nutrients back to the plot. These plants are so good at up-taking nutrients from the soil and making them readily available for your deer herd through the leaves, that you must be willing to soil test and add back nutrients to stay at a high level of nutrition. This is really easy, but its the secret to continuously having extremely productive food plots. There you have it. The blue print to a successful plot.

1. Wild Turkey Biology

Wild turkeys, Meleagris gallopavo, belong to the Galliformes family of birds which also include grouse, pheasants, and peacocks. Wild turkeys found in New Brunswick are Eastern wild turkeys, one of 5 distinct subspecies of turkey that inhabit North America.


Wild turkeys range in weight from 2.7 to 13.6 kg (6 to 30 lbs) and may be more than a metre (40 inches) in height when displaying. Adult male birds are much larger than females, are generally darker in colour with a bare head and neck and feature a “beard” of feathers growing from the center of their chest. Approximately 5,000 to 6,000 feathers cover a wild turkey’s body. The plumage of male wild turkeys is iridescent with metallic green, copper and bronze colours. Female wild turkeys are a dull brown and grey in colour.

Map showing wild turkey distribution in Canada and provinces that have a turkey hunting season. Source: Canadian Wild Turkey Federation.


New Brunswick wild turkeys are found primarily along the province’s western border with the state of Maine and in the southwestern corner of the province.

Although New Brunswick is near the northeastern limit of the Eastern wild turkey, populations in the province are a result of introductions. Other Canadian provinces that have wild turkey populations and hunting seasons include British Columbia, Alberta, Manitoba, Ontario, and Quebec.

New Brunswick’s wild turkey population is currently estimated at 2,000 to 3,000 birds.


An adult male turkey is called a “tom” or “gobbler” while an adult female is called a “hen”. A one-year old juvenile wild turkey is referred to as a “jake” if a male and a “jenny” if female.


The average life span of a wild turkey is 2.5 years although some birds have been known to live up to 5 years.


Wild turkeys are primarily ground foragers and eat a variety of plants, insects and invertebrates. Plant foods include acorns, leaves, buds, grasses, seeds and waste grains. Other foods include beetles, snails, grasshoppers, frogs and small snakes. Young turkeys rely heavily on protein rich insects.Wild turkeys will scratch through debris on the ground to uncover food as well as pluck it directly off plants.

Wild turkeys are primarily ground feeders and eat a variety
of plant, insect, and invertebrate foods.


Eastern wild turkeys prefer hardwood and mixed conifer-hardwood forests with scattered openings such as pastures and fields.


Wild turkeys are social and will live in flocks of 10 or more birds especially during the winter. Flocks break into smaller groups prior to spring breeding season. During the daytime wild turkeys spend most of their time on the ground while at night they roost in trees to avoid predators. Turkeys will usually run rather than fly when they sense danger. They are capable of ground speeds of 40 km/hr (25 mph) and flight speeds up to 80 km/hr (50 mph) for short distances.

Although they do not migrate, wild turkeys may travel some distance from their breeding site especially during the winter to locate food sources.


During breeding season from March to May, male turkeys display and gobble to attract females and assert dominance over subordinate males. Males are polygamous and will mate with several hens.

Following mating, hens lay 9 to 13 eggs in a shallow ground nest over a period of about two weeks. Once the eggs are laid the hen will incubate them for approximately 4 weeks. The creamy white to light brown speckled eggs are slightly larger than a hen’s egg.

Newborn turkeys or “poults” are fully feathered and able to leave the nest within 12 to 24 hours of hatching to forage for food with their mother. Typically, more than 50% of the poults do not survive due to weather and predation. Adult male turkeys do not play a role in incubating the eggs or raising the young.

Newborn turkeys or “poults” are able to leave the nest
within 12 to 24 hours of hatching.


Wild Turkeys have keen eyesight and a strong ability to sense movement. Their eyes are located on opposite sides of their head giving them a stationary field of view that exceeds 300 0 . They also distinguish colours very well.

The turkey’s hearing ability is also well developed despite the fact they lack an external ear structure and rely on small openings located behind their eyes to hear. Studies indicate turkeys hear both above and below the frequency range heard by humans and are adept at locating distant sounds.

Turkeys do not have a strong sense of smell. The region of a turkey’s brain that controls smells is small in comparison to other animals.


The most distinctive call of the wild turkey is the “gobble-gobble-gobble” call used by males during the spring breeding season. Other calls include “clucks”, “purrs” “yelps” and “putts”. Turkeys call to attract mates, signal danger, or keep in touch with the rest of the flock.

Wild turkeys possess keen eyesight
and well developed hearing.

Examples of turkey calls:

Audio clips provided by Gulvas Wildlife Adventures, courtesy of the National Wildlife Turkey Federation

Gobbles are the one of the main vocalizations of male turkeys and are used to attract hens during the spring mating season. “Gobbling” is not a recommended turkey call since it may attract other hunters to your location or scare away less dominant male birds.

Audio clip

Clucks are used by one turkey to get the attention of another. This type of call is used by hunters to reassure an approaching gobbler that a hen is waiting for it.

Audio clip

Purrs are sounds made by turkeys when they are content and are often made by feeding turkeys. Purr vocalizations allow turkeys to maintain contact with one another.

Audio clip

Yelps are one of the most commonly heard turkey vocalizations and are made by both male and female turkeys. Yelps are used by hens to communicate with gobblers during the spring mating season and are an important hunting call for turkey hunters.

Audio clip

Putts are the turkey’s alarm call and are used to signal danger.

Audio clip

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Native Species That Can Resemble Turkey Berry

Control and management recommendations vary according to individual circumstances. Location, habitat, weather, and a variety of other conditions are factors that help determine the best treatment choice. To find the safest and most effective treatment for your situation, consult your state’s land-grant institution. If you will use chemicals as part of the control process, always refer to the product label.

United States Land-Grant University System – Find your land-grant university’s college of agriculture, Cooperative Extension office, or other related partner on this map provided by USDA.

Images and Information – University of Florida – Center for Aquatic and Invasive Plants

Rio Grande Wild Turkey

The Rio Grande wild turkey (Meleagris gallopavo intermedia) has the largest population and the widest range of the three turkey subspecies (Rio Grande, Merriam’s, and Eastern wild turkeys) found in Texas. Unregulated hunting in the 1800s greatly reduced the Rio Grande wild turkey (RGWT) population in Texas to about 100,000 birds by 1920. Since then, their numbers have recovered thanks to better habitat management, restocking programs by Texas Parks and Wildlife Department (TPWD), and partnerships with landowners and conservation groups. However, there has been a steady decline in their populations in certain regions since the 1970s, which prompted TPWD to partner with universities to study the biology and habitat requirements of the turkeys in different parts of the states.

The RGWT do not occupy the Trans Pecos and High Plains ecoregions of Texas. Typically, they inhabit areas that receive enough rainfall to sustain their food sources. The Edwards Plateau is their historic geographic center and has the highest number of RGWT today. Male turkeys are referred to as “toms” males of breeding age are “gobblers” females are “hens” juveniles are “poults.”

Physical Attributes

Male Rio Grand Wild Turkey (Gobbler)

The bodies of wild turkeys are covered with 5,000 to 6,000 feathers. These feathers provide insulation, lift during flight, and touch sensation and ornamentation. A wild turkey undergoes five molts (feather replacement) during its lifetime: natal, juvenile, first basic, alternate (first winter), and basic (adult plumage). The body feathers of toms are vibrantly colored: iridescent copper, bronze, red, green, and gold. While hens have these same colors, they are significantly dulled and muted which causes the female to appear brown. Gobblers have a beard, which is a cluster of long follicles in the center of their chest that can be an inch to 10 inches long. Unlike the rest of the body feathers which undergo 5 molts throughout an RGWT’s lifetime, the beard does not molt. It becomes visible when the turkey is 6-7 months of age, and it continues to grow throughout their lifetime. Hens may occasionally have beards, although this uncommon event produces much shorter beards than toms. Both toms and hens have sparsely feathered heads with bare legs and feet that are pink to red in color. Toms grow a spur on the lower third of their leg that starts off small and rounded, but which becomes pointed and about 2 inches long with time. Hens also have a spur, although their spurs stay small and blunted. Gobblers weigh 17-21 pounds and attain a height of 40 inches. Hens weigh 8-11 pounds and are 30 inches tall. Poults are relatively small when hatched they only weigh 2 ounces.

Mating and nesting

Gobblers attract a hen’s attention by gobbling and strutting.

Gobblers attract a hen’s attention by gobbling and strutting. The hen will select a mate by lying close to the ground in front of him, which signals the male to begin copulation. Nesting begins in early spring in south Texas and continues on through July and August in central and north Texas. Hens choose nesting sites that are in grass clumps, brush piles, understory brush, or leaf litter.

Rio Grande Wild Turkey Nest

They scratch out a shallow depression or bowl shape in these selected areas to form the nest. In order to protect their nests, hens choose sites that have enough herbaceous cover to hide the poults while still allowing the hen an unobstructed view to watch for threats. Nest sites are typically found in areas that are within ¼ mile of a water source, have grass heights of 18 inches, and have a high abundance of insects. Females lay one egg per day, producing an average total of 10-11 eggs. The eggs are cream or tan in color and may have brown speckles. Incubation lasts around 28 days after the last egg is laid. After hatching, the hens and poults form a brood this brood forages together for the majority of the day. At this young age, poults mainly consume invertebrates such as grasshoppers, beetles, and spiders. The poults continue to roost on the ground for the first two weeks of life until their natal plumage is replaced by flight feathers and they can fly up into roost trees.

Habitat and diet

The Rio Grande wild turkey is an opportunistic forager that feeds on green foliage, insects, seeds from grasses and forbs, and mast (acorns and nuts). Annually, their diet consists of about 36% grasses, 29% insects, 19% mast, and 16% forbs. The individual plant species that a turkey uses as food varies by region and by season. Water is very important to RGWTs this includes surface water, metabolic water, and preformed water. Surface water is free-standing water such as ponds, creeks, or troughs. Metabolic water is derived from foods when they are digested. Preformed water is contained within the food itself, such as the water found in succulent plants.

RGWTs require a habitat that contains an interspersion of wooded and open areas. A mix of these areas, while individually important to the turkeys, also provides the added benefit of large amounts of edge habitat. Edge habitat is useful for escaping from both predators and the heat. Although RGWTs do not migrate, they have two distinct roosting sites: summer roosts and winter roosts. They prefer to roost in tall hardwood trees that have broad canopies with many horizontal limbs. Species such as live oak, hackberry, pecan, elm, and cottonwood exhibit these traits. In areas that lack trees, artificial roosts can provide similar benefits. RGWTs prefer to roost over an open understory, which is why they can often be seen roosting over water. Roosting areas consist of 10-15 acres with a 50-70% canopy cover. An important part of the roosting area is the area that surrounds it turkeys need open spaces within 100 yards of roost trees to serve as launching and landing pads for ascending and descending from the roosts. In general, these highly mobile birds need large amounts of usable space, as they annually travel between 6 and 26 miles. For example, their range in the Rolling Plains varies between 2,400 and 5,900 acres, whereas their range in the Edwards Plateau varies from 3,800 to 6,600 acres. In the spring, bred hens move independently from hens that did not breed. In the summer, gobblers separate from juvenile males and non-breeding females. In the late summer, brood flocks form. In the winter, the males join the flocks.

Predation and other mortality factors

Foxes are a common nest predator

Because hens nest on the ground, the reproductive success of RGWTs depends on weather and yearly rainfall, the range condition, and the body condition of the individual hen. In addition to these factors, there are many predators that prey on the nests and hens. The most common of these nest predators are raccoons and foxes although reptiles, owls, hawks, skunks, feral hogs, and other mammals will also prey on the hens, eggs, and poults. Several different predators may depredate the same nest, but they do not always consume the entire clutch of eggs. In some instances, hens may resume incubating the surviving eggs after depredation of the nest. Life does not get much easier for poults after hatching, as many of the same species that prey on nests also prey on poults. On average, there is a 60-75% nest failure and a 12-50% poult survival rate annually. Survival increases once they begin roosting in trees rather than on the ground.

Wild turkeys are susceptible to a number of diseases. These diseases include ones that are common among domestic poultry, such as mycoplasmosis, salmonellosis, aspergillosis, and reticuloendothelosis, which affect the immune system and causes abnormal internal growth. The avian pox causes lesions which prevents the affected turkey from foraging and makes them more prone to predation.

Conservation and management

The decline of RGWTs in certain parts of their range may be due to these predation and mortality factors, in addition to changes in land use, land fragmentation, and an increase in brush height/density. Interested landowners and land managers can manage their land for turkey in multiple ways: preventing cattle from overgrazing, conducting prescribed burns, and practicing brush control, among others. Ensuring sufficient nesting and brooding habitat will increase nesting success and poult survival resulting in more individuals being recruited into the population, and maintaining roost sites will enhance the usability of the land even more.

For more information about the biology and habitat requirement of RGWT and general recommendations for landowners to implement, see the publications that AgriLife has developed below.

ID large-thorned plant growing in Turkey - Biology

Scientific name: Polystichum acrostichoides
Common name: Christmas fern

(Information in this species page was compiled in part by Jon Belasco as part of the Biology 220W class at Penn State New Kensington in Spring 2001)

Polystichum acrostichoides is called the "Christmas fern" because some parts of the plant remain green throughout the year and are thus available for use in decorations at Christmas time. The rich, green leaves (fronds) of the fern are up to three feet long and are about four inches wide. They are tough, leathery and lance-shaped to a pointed tip. The fronds are attached to a relatively short stalk that is brown at its soil base and green toward its apex. A central, perennial root (the rhizome) supports the stalk that then supports the arching clump of tall fronds. The sterile fronds (fronds not involved in reproduction) are shorter and less well supported than the spore producing fertile fronds. The sterile fronds, then, form an encircling, arching border around the central mass of taller, more erectly held fertile fronds. It is the sterile fronds which remain green throughout the year.

Each fertile frond sports 20-40 leaflets, with the sori (clusters of spore-producing sporangia) forming only on the upper third to half of the leaflets. These fertile leaflets are noticeably shorter than the non-sori bearing leaflets, giving the impression of a sudden narrowing of the entire fertile frond toward the top.

In a typical fern life cycle, spores are produced within sporangia when moisture and temperature conditions are favorable. In the Christmas fern, spore production occurs between June and October. A small percentage of the released spores from the tall, erectly held fertile fronds will find an optimal microhabitat in the surrounding soil and then be able to grow into the fern's tiny sperm and ova producing life stage called the gametophyte. Fusion of the sperm and ova within the gametophyte will produce a new sporophyte which can then grow into a mature fern.

New sporophytes can also arise vegetatively from the perennial rhizome. These coiled young ferns are called fiddleheads (see image at right). Fiddleheads arise in the early to mid-spring.

Christmas fern is found in the north-eastern and north-central portions of North America from New Brunswick south to North Carolina. It is especially abundant on well shaded, forested hill sides near streams. Soils of moderate moisture and a more neutral pH are preferred. This pH preference is reflected in the increased densities of Christmas ferns in soils that overlie limestone bedrock. Christmas fern is seldom found in soils that are too waterlogged or that are too rocky.

Ecological Significance
Christmas fern may grow in large, extensive colonial masses but more typically is found in clusters of two or three individuals. Growing ferns and the accumulated detritus of past sterile fronds form a dense covering mass over the soil surface. This mass helps to stabilize the underlying soil and prevent or lessen erosion. It also generates a protective, concealing habitat for a number of ground feeding and ground nesting bird species.

Because of their complex chemical composition, ferns are eaten by very few browsers or grazers. Fern densities in general have been increasing in many of the forests of Pennsylvania due to the removal of competing, but more palatable, under-story plant species by the extensive browsing by the state's rapidly increasing deer population. This change in the forest under-story habitat structure has had wide-spread and deleterious impacts on the densities and successes of some mid-canopy birds (like the least flycatcher and the yellow billed cuckoo). Other bird species (like the wild turkey, for example) that use the concealing fern masses for their nesting sites have, however, increased in density in these "fern park" forests.

In commercially grown Christmas ferns a fern scale insect and the mealy bug can extensively feed on and damage the plants. Also, the larvae of the pyralid moth (Herperogramma aeglealis) can construct a variety of feeding shelters within the sterile fronds of the Christmas fern and consume its terminal frond leaflets (probably before they accumulate significant levels of their protective chemicals).

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V. thapsus is a dicotyledonous plant that produces a rosette of leaves in its first year of growth. [3] [4] The leaves are large, up to 50 cm long. The second-year plants normally produce a single unbranched stem, usually 1–2 m tall. In the eastern part of its range in China, it is, however, only reported to grow up to 1.5 m tall. [5] The tall, pole-like stems end in a dense spike of flowers [3] that can occupy up to half the stem length. All parts of the plants are covered with star-shaped trichomes. [5] [6] This cover is particularly thick on the leaves, giving them a silvery appearance. The species' chromosome number is 2n = 36. [7]

On flowering plants, the leaves are alternately arranged up the stem. They are thick and decurrent, with much variation in leaf shape between the upper and lower leaves on the stem, ranging from oblong to oblanceolate, and reaching sizes up to 50 cm long and 14 cm across (19 inches long and 5 inches wide). [8] [9] They become smaller higher up the stem, [3] [4] and less strongly decurrent down the stem. [3] The flowering stem is solid and 2–2.5 cm (nearly an inch) across, and occasionally branched just below the inflorescence, [4] usually following damage. [10] After flowering and seed release, the stem and fruits usually persist in winter, [11] drying into dark brown, stiff structures of densely packed, ovoid-shaped, and dry seed capsules. The dried stems may persist into the following spring or even the next summer. The plant produces a shallow taproot. [9]

Flowers are pentamerous with (usually) five stamen, a five-lobed calyx tube and a five-petalled corolla, the latter bright yellow and an 1.5–3 cm (0.59–1.18 in) wide. The flowers are almost sessile, with very short pedicels (2 mm, 0.08 in). The five stamens are of two types, with the three upper stamens being shorter, their filaments covered by yellow or whitish hairs, and having smaller anthers, while the lower two stamens have glabrous filaments and larger anthers. [6] [note 1] The plant produces small, ovoid (6 mm, 0.24 in) capsules that split open by way of two valves, each capsule containing large numbers of minute, brown seeds less than 1 mm (0.04 in) [12] in size, marked with longitudinal ridges. A white-flowered form, V. thapsus f. candicans, is known to occur. [13] Flowering lasts up to three months from early to late summer (June to August in northern Europe), [4] with flowering starting at the bottom of the spike and progressing irregularly upward each flower opens for part of a day and only a few open at the same time around the stem. [11]

For the purpose of botanical nomenclature, Verbascum thapsus was first described by Carl Linnaeus in his 1753 Species Plantarum. The specific epithet thapsus had been first used by Theophrastus (as Θάψος , Thapsos) [14] for an unspecified herb from the Ancient Greek settlement of Thapsos, near modern Syracuse, Sicily, [14] [15] though it is often assimilated to the ancient Tunisian city of Thapsus. [16]

At the time, no type specimen was specified, as the practice only arose later, in the 19th century. When a lectotype (type selected amongst original material) was designated, it was assigned to specimen 242.1 of Linnaeus' herbarium, the only V. thapsus specimen. [note 2] The species had previously been designated as type species for Verbascum. [18] European plants exhibit considerable phenotypical variation, [19] which has led to the plant acquiring many synonyms over the years. [17] [20] Introduced American populations show much less variation. [19]

The taxonomy of Verbascum has not undergone any significant revision since Svanve Mürbeck's monographies in the 1930s, with the exception of the work of Arthur Huber-Morath, who used informal group in organizing the genus for the florae of Iran and Turkey to account for many intermediate species. Since Huber-Morath's groups are not taxonomical, Mürbeck's treatment is the most current one available, as no study has yet sought to apply genetic or molecular data extensively to the genus. In Mürbeck's classification, V. thapsus is placed in section Bothrospermae subsect. Fasciculata (or sect. Verbascum subsect. Verbascum depending on nomenclatural choices) alongside species such as Verbascum nigrum (black or dark mullein), Verbascum lychnitis (white mullein) and Verbascum sinuatum (wavy-leaved mullein). [21] [22] [23] [24]

Subspecies and hybrids Edit

Hybrids of Verbascum thapsus [7] [25]
Hybrid name Other
parent species
V. × duernsteinense Teyber V. speciosum
V. × godronii Boreau V. pulverulentum
V. × kerneri Fritsch V. phlomoides
V. × lemaitrei Boreau V. virgatum
V. × pterocaulon Franch. V. blattaria
V. × thapsi L. V. lychnitis syn. V. × spurium
W.D.J.Koch, may be a
nomen ambiguum [26]
V. × semialbum Chaub. V. nigrum
none V. pyramidatum

There are three usually recognized subspecies:

  • V. thapsus subsp. thapsus type, widespread.
  • V. thapsus subsp. crassifolium (Lam.) Murb. Mediterranean region and to 2000 metres in southwestern Austria. [27] (syn. subsp. montanum (Scrad.) Bonnier & Layens)
  • V. thapsus subsp. giganteum (Willk.) Nyman Spain, endemic.

In all subspecies but the type, the lower stamens are also hairy. [28] In subsp. crassifolium, the hairiness is less dense and often absent from the upper part of the anthers, while lower leaves are hardly decurrent and have longer petioles. [27] In subsp. giganteum, the hairs are densely white tomentose, and lower leaves strongly decurrent. Subsp. crassifolium also differs from the type in having slightly larger flowers, which measure 15–30 mm wide, whereas in the type they are 12–20 mm in diameter. [27] Both subsp. giganteum and subsp. crassifolium were originally described as species. [3] Due to its morphological variation, V. thapsus has had a great many subspecies described. A recent revision led its author to maintain V. giganteum but sink V. crassifolium into synonymy. [24]

The plant is also parent to several hybrids (see table). Of these, the most common is V. × semialbum Chaub. (× V. nigrum). [7] All occur in Eurasia, [7] and three, V. × kerneri Fritsch, V. × pterocaulon Franch. and V. × thapsi L. (syn. V. × spurium W.D.J.Koch), have also been reported in North America. [25] [29]

Common names Edit

V. thapsus is known by a variety of names. European reference books call it "great mullein". [30] [31] [32] In North America, "common mullein" is used [33] [34] while western United States residents commonly refer to mullein as "cowboy toilet paper". [35] [36]

In the 19th century it had well over 40 different common names in English alone. Some of the more whimsical ones included "hig candlewick", "indian rag weed", "bullicks lungwort", "Adams-rod", "hare's-beard" and "ice-leaf". [37] Vernacular names include innumerable references to the plant's hairiness: "woolly mullein", "velvet mullein" or "blanket mullein", [32] [38] "beggar's blanket", "Moses' blanket", "poor man's blanket", "Our Lady's blanket" or "old man's blanket", [31] [34] [39] and "feltwort", and so on ("flannel" is another common generic name). "Mullein" itself derives from the French word for "soft". [40]

Some names refer to the plant's size and shape: "shepherd's club(s)" or "staff", "Aaron's Rod" (a name it shares with a number of other plants with tall, yellow inflorescences), and a plethora of other "X's staff" and "X's rod". [31] [34] [41] The name "velvet dock" or "mullein dock" is also recorded, where "dock" is a British name applied to any broad-leaved plant. [42]

Verbascum thapsus has a wide native range including Europe, northern Africa and Asia, from the Azores and Canary Islands east to western China, north to the British Isles, Scandinavia and Siberia, and south to the Himalayas. [5] [43] [44] In northern Europe, it grows from sea level up to 1,850 m altitude, [4] while in China it grows at 1,400–3,200 m altitude. [5]

It has been introduced throughout the temperate world, and is established as a weed in Australia, New Zealand, tropical Asia, La Réunion, North America, Hawaii, Chile, Hispaniola and Argentina. [44] [45] [46] [47] It has also been reported in Japan. [48]

In the United States it was imported very early in the 18th [note 3] century and cultivated for its medicinal and piscicide properties. By 1818, it had begun spreading so much that Amos Eaton thought it was a native plant. [note 4] [9] [49] In 1839 it was already reported in Michigan and in 1876, in California. [9] It is now found commonly in all the states. [50] In Canada, it is most common in the Maritime Provinces as well as southern Quebec, Ontario and British Columbia, with scattered populations in between. [19] [51]

Great mullein most frequently grows as a colonist of bare and disturbed soil, usually on sandy or chalky ones. [7] It grows best in dry, sandy or gravelly soils, although it can grow in a variety of habitats, including banksides, meadows, roadsides, forest clearings and pastures. This ability to grow in a wide range of habitats has been linked to strong phenotype variation rather than adaptation capacities. [52]

Great mullein is a biennial and generally requires winter dormancy before it can flower. [10] This dormancy is linked to starch degradation activated by low temperatures in the root, and gibberellin application bypasses this requirement. [53] Seeds germinate almost solely in bare soil, at temperatures between 10 °C and 40 °C. [10] While they can germinate in total darkness if proper conditions are present (tests give a 35% germination rate under ideal conditions), in the wild, they in practice only do so when exposed to light, or very close to the soil surface, which explains the plant's habitat preferences. While it can also grow in areas where some vegetation already exists, growth of the rosettes on bare soil is four to seven times more rapid. [10]

Seeds germinate in spring and summer. Those that germinate in autumn produce plants that overwinter if they are large enough, while rosettes less than 15 cm (6 in) across die in winter. After flowering the entire plant usually dies at the end of its second year, [10] but some individuals, especially in the northern parts of the range, require a longer growth period and flower in their third year. Under better growing conditions, some individuals flower in the first year. [54] Triennial individuals have been found to produce fewer seeds than biennial and annual ones. While year of flowering and size are linked to the environment, most other characteristics appear to be genetic. [55]

A given flower is open only for a single day, opening before dawn and closing in the afternoon. [19] Flowers are self-fecundating and protogynous (with female parts maturing first), [19] and will self-pollinate if they have not been pollinated by insects during the day. While many insects visit the flowers, only some bees actually accomplish pollination. The flowering period of V. thapsus lasts from June to August in most of its range, extending to September or October in warmer climates. [9] [10] [12] Visitors include halictid bees and hoverflies. [11] The hair on lower stamens may serve to provide footholds for visitors. [19]

The seeds maintain their germinative powers for decades, up to a hundred years, according to some studies. [56] Because of this, and because the plant is an extremely prolific seed bearer (each plant produces hundreds of capsules, each containing up to 700+ seeds, [19] with a total up to 180,000 [9] [10] or 240,000 [12] seeds), it remains in the soil seed bank for extended periods of time, and can sprout from apparently bare ground, [10] or shortly after forest fires long after previous plants have died. [12] Its population pattern typically consists of an ephemeral adult population followed by a long period of dormancy as seeds. [19] Great mullein rarely establishes on new grounds without human intervention because its seeds do not disperse very far. Seed dispersion requires the stem to be moved by wind or animal movement 75% of the seeds fall within 1 m of the parent plant, and 93% fall within 5 m. [10]

Megachilid bees of the genus Anthidium use the hair (amongst that of various woolly plants) in making their nests. [57] The seeds are generally too small for birds to feed on, [11] although the American goldfinch has been reported to consume them. [58] Other bird species have been reported to consume the leaves (Hawaiian goose) [59] or flowers (palila), [60] or to use the plant as a source when foraging for insects (white-headed woodpecker). [61] Additionally, deer and elk eat the leaves. [62]

Seed of Verbascum thapsus has been recorded from part of the Cromer Forest Bed series and at West Wittering in Sussex from some parts of the Ipswichian interglacial layers. [63]

Because it cannot compete with established plants, great mullein is no longer considered a serious agricultural weed and is easily crowded out in cultivation, [19] except in areas where vegetation is sparse to begin with, such as Californian semi-desertic areas of the eastern Sierra Nevada. In such ecological contexts, it crowds out native herbs and grasses its tendency to appear after forest fires also disturbs the normal ecological succession. [10] [12] Although not an agricultural threat, its presence can be very difficult completely to eradicate and is especially problematic in overgrazed pastures. [9] [10] [12] The species is legally listed as a noxious weed in the American state of Colorado (Class C) [64] and Hawaii, [65] and the Australian state of Victoria (regionally prohibited in the West Gippsland region, and regionally controlled in several others). [66]

Despite not being an agricultural weed in itself, it hosts a number of insects and diseases, including both pests and beneficial insects. [67] It is also a potential reservoir of the cucumber mosaic virus, Erysiphum cichoraceum (the cucurbit powdery mildew) and Texas root rot. [19] [68] A study found V. thapsus hosts insects from 29 different families. Most of the pests found were western flower thrips (Frankliniella occidentalis), Lygus species such as the tarnished plant bug (L. lineolaris), and various spider mites from the family Tetranychidae. These make the plant a potential reservoir for overwintering pests. [67]

Other insects commonly found on great mullein feed exclusively on Verbascum species in general or V. thapsus in particular. They include mullein thrips (Haplothrips verbasci), [67] Gymnaetron tetrum (whose larva consume the seeds) and the mullein moth (Cucullia verbasci). [9] Useful insects are also hosted by great mullein, including predatory mites of the genera Galendromus, Typhlodromus and Amblyseius, the minute pirate bug Orius tristicolor [67] and the mullein plant bug (Campylomma verbasci). [69] The plant's ability to host both pests and beneficials makes it potentially useful to maintain stable populations of insects used for biological control in other cultures, like Campylomma verbasci and Dicyphus hesperus (Miridae), a predator of whiteflies. [70] [71] A number of pest Lepidoptera species, including the stalk borer (Papaipema nebris) and gray hairstreak (Strymon melinus), also use V. thapsus as a host plant. [72]

Control of the plant, when desired, is best managed via mechanical means, such as hand pulling and hoeing, preferably followed by sowing of native plants. Animals rarely graze it because of its irritating hairs, and liquid herbicides require surfactants to be effective, as the hair causes water to roll off the plant, much like the lotus effect. Burning is ineffective, as it only creates new bare areas for seedlings to occupy. [9] [10] [12] G. tetrum and Cucullia verbasci usually have little effect on V. thapsus populations as a whole. [12] Goats and chickens have also been proposed to control mullein. [10] Effective (when used with a surfactant) contact herbicides include glyphosate, [9] [12] triclopyr [9] and sulfurometuron-methyl. [12] Ground herbicides, like tebuthiuron, are also effective, but recreate bare ground and require repeated application to prevent regrowth. [10]

Phytochemicals Edit

Phytochemicals in Verbascum thapsus flowers and leaves include saponins, polysaccharides, mucilage, flavonoids, tannins, iridoid and lignin glycosides, and essential oils. [2] The plant's leaves, in addition to the seeds, have been reported to contain rotenone, although quantities are unknown. [73]

Traditional medicine Edit

Although long used in herbal medicine, there are no drugs manufactured from its components. [2] Dioscorides first recommended the plant 2000 years ago, believing it useful as a folk medicine for pulmonary diseases. [74] Leaves were smoked to attempt to treat lung ailments, a tradition that in America was rapidly transmitted to Native American peoples. [31] [75] The Zuni people, however, use the plant in poultices of powdered root applied to sores, rashes and skin infections. An infusion of the root is also used to treat athlete's foot. [76] All preparations meant to be drunk have to be finely filtered to eliminate the irritating hairs. [53]

Oil from the flowers was used against catarrhs, colics, earaches, frostbite, eczema and other external conditions. [31] Topical application of various V. thapsus-based preparations was recommended for the treatment of warts, [77] boils, carbuncles, hemorrhoids, and chilblains, amongst others. [31] [75] Glycyrrhizin compounds with bactericide effects in vitro were isolated from flowers. [78] The German Commission E describes uses of the plant for respiratory infections. [79] It was also part of the National Formulary in the United States [75] and United Kingdom. [31]

The plant has been used in an attempt to treat colds, croup, sunburn and other skin irritations. [80]

Other uses Edit

Roman soldiers are said to have dipped the plant stalks in grease for use as torches. Other cultures use the leaves as wicks. [80] Native Americans and American colonists lined their shoes with leaves from the plant to keep out the cold. [80] [31] [75]

Mullein may be cultivated as an ornamental plant. [1] As for many plants, (Pliny the Elder described it in his Naturalis Historia), [note 5] great mullein was linked to witches, [31] although the relationship remained generally ambiguous, and the plant was also widely held to ward off curses and evil spirits. [31] [53] [74] [75] The seeds contain several compounds (saponins, glycosides, coumarin, rotenone) that are toxic to fish, and have been widely used as piscicide for fishing. [9] [82]

Due to its weedy capacities, the plant, unlike other species of the genus (such as V. phoeniceum), is not often cultivated. [1]

  1. ^ They are all hairy in subspecies V. crassifolium and V. giganteum.
  2. ^ The lectotypification is usually attributed to Arthur Huber-Morath (1971) Denkschriften der Schweizerischen Naturforschenden Gesellschaft 87:43. Some disagree since Huber-Morath did not specifically cite sheet 242.1, and credit instead L. H. Cramer, in Dassanayake & Fosberg (1981) A Revised Handbook to the Flora of Ceylon 3:389. [17]
  3. ^ The 1630 number in Mitch may be a typo: the beginning of the 18th century is cited in other sources. [9][19]
  4. ^ Eaton went so far as to write: "When botanists are so infatuated with wild speculation, as to tell us the mullein was introduced, they give our youngest pupils occasion to sneer at their teachers." [13]
  5. ^ In book 25, Pliny describes "two principal kinds [of verbascum]" thought to be V. thapsus and V. sinuatum. The precise attribution of a third kind is unclear. [81]
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Bibliography Edit

  • Watts, Donald (2000). Elsevier's Dictionary of Plant Names and their Origin. Amsterdam: Elsevier Science. ISBN0-444-50356-0 .

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The leaves of these plants are the most obvious difference. The stiff leaf stems, or petioles, of Alocasia extend into the leaves. This causes the leaves to follow the line of the petioles. As a result, most Alocasia leaves tend to point upwards. Some Alocasia varieties have leaves that extend horizontally. By contrast, the petioles of Colocasia connect down from the notches in the leaves. This enables the leaves to droop or hang at a downward angle. This visual distinction makes it easy to tell these plants apart: If the leaves point up, the plant is probably an Alocasia if the leaves point down, it’s probably a Colocasia.


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Plant materials and sample preparation for analysis

Roots of Rh. tanguticum plants were collected from Qinghai, Gansu, and Sichuan provinces. Under canopy and open habitats were selected as the sampling sites in each province where five whole Rh. tanguticum plants were excavated from each site for further analysis (Fig. 8). Samples were collected from August to September in a period ranging from 2016 to 2018 with the date of collection and geographical information being shown in Table 1. Plant height, root length, root diameter, leaf length, length of leaf lobes, and percentage leaf lobes (leaf lobes length to leaf length ratio) were measured after excavation of the plants. All samples were identified by Professor Guoying Zhou, and the roots were washed and air dried. The specimens were then stored at the herbarium in Northwest Institute of Plateau Biology, Chinese Academy of Science.

Rh. tanguticum in the (a) under canopy habitats and (b) open habitats

Chemicals and reagents

Chemicals and solvents used in the study are all of HPLC grade. Water, acetonitrile, methanol, and formic acid were purchased from CNW Technologies GmbH (Düsseldorf, Germany), while L-2-chlorophenylalanine was purchased from Shanghai Hengchuang Bio-technology Co., Ltd. (Shanghai, China). Reference standards of emodin, aloe-emodin, rhein, chrysophanol, physcion, sennoside A, and sennoside B were used for contents calculate based on the methods described by our previous study [66].

Soil properties

The soil humidity was measured using a portable TDR-100 soil moisture probe (Spectrum Technologies, Inc., Plainfield, IL, USA). Soil samples were collected alongside the plant roots at a depth of 0–10 cm. Air-dried soil samples were sieved through a 2-mm mesh and then ground to fine powder. The total nitrogen content of the dry soil samples was determined using the semimicro-Kjeldahl method [67] while the soil organic carbon content was determined using the Walkley-Black method [68].

Untargeted UPLC-MS extraction and analysis

80 mg of the plant sample powder was weighed and introduced into a 1 mL solvent containing methanol and water (7:3 V/V ratio). This was followed by the addition of 20 μL of L-2-chlorophenylalanine (0.3 mg/mL) in methanol, and 20 μL of C-17 (0.01 mg/mL) in methanol which acted as the internal standard. Two small steel balls were then added followed by pre-cooling at − 20 °C for 2 min before they were put into an automatic sample rapid grinder (JXFSTPRP-24/32, Shanghai, China) at 60 Hz for 2 min. The samples were then placed in an ultrasonic cleaner (SB-5200DT, Ningbo, Zhejiang, China) for 30 min followed by cooling at − 20 °C for 20 min. The samples were then centrifuged at 13000 rpm (TGL-16MS, Shanghai, China) for 10 min at 4 °C. A syringe was used to suck out 200 μL of the supernatant, and the supernatant was then filtered through a 0.22 μm organic membrane into an LC sample injection vial and stored at − 80 °C. All the reagents used for the extraction were pre-cooled at − 20 °C before use. On the other hand, all samples were mixed to a quality control (QC) sample using the same volume, and all the QC samples had the same volume as the samples used.

UPLC-QTOF-MS E analysis was performed using an ACQUITY UPLC system (Waters Corporation, Milford, MA, USA) and a Waters Xevo G2-XS QTof mass spectrometer System (Waters Corporation, Milford, MA, USA) coupled with an electrospray ionization (ESI) interface where the results were detected in both ESI positive and ESI negative ion modes. The UPLC system was equipped with an ACQUITY UPLC BEH C18 column (1.7 μm, 2.1 × 100 mm). The mobile phase composed of mobile A (0.1% formic acid in water, v/v) and mobile B (0.1% formic acid in acetonitrile, v/v). The elution condition was set as: 0 min, 1% B 1 min, 5% B 2 min, 25% B 30 min, 60% B 3.5 min, 60% B 7.5 min, 90% B 9.5 min, 100% B 12.5 min, 100% B 12.7 min, 1% B, and 16 min, 1% B. The flow rate was set at 0.4 mL/min while the column temperature was 45 °C. The injection volume was 2 μL and all samples were kept at 4 °C throughout the analysis.

The spectrum conditions were: source temperature, 120 °C source offset, 80 V reference capillary voltage, 2.5 kV cone voltage, 40 V desolvation gas temperature, 450 °C desolvation gas flow, 800 L/h and cone gas flow, 50 L/h. Additionally, the capillary voltage for the negative mode was -2 kV while the capillary voltage for the positive mode was + 3 kV. The collision dissociation gas used was argon (99.999%) while the desolvation and cone gas was nitrogen (> 99.5%). The data was acquired from 50 m/z to 1000 m/z in the MS E centroid full scan mode by rapidly switching from a low energy (CE 6 eV) scan to a high energy (CE ramp 20-35 eV) scan with the scan rate being 0.1 s/scan. The external reference for lock mass correction was infused at a flow of 5 μL/min through the reference probe for 30s each. Leucine- enkephalin, performed as acetonitrile/water/formic acid (50:49.9:0.1, v/ v/v) in 250 ng/mL standard solution, was used as the external reference. The QC samples were injected for every 10 samples to enhance the data repeatability.

Untargeted UPLC data processing

The raw data obtained from UPLC-MS was collected using the UNIFI 1.8.1 (Waters Corporation, Milford, MA, USA) and then imported to the Progenesis QI software (Nonlinear Dynamics, Newcastle, UK). The parameters were set as: 5 ppm for precursor tolerance, 10 ppm for fragment tolerance, and 0.02 min as the retention time (RT) tolerance. In addition, the internal standard detection parameters were employed for perk RT alignment while the noise level and minimum intensity were set at 10.00 and 15% of base peak intensity respectively to eliminate noise peaks. As a result, a 3D matrix expressed with m/z, peak RT, and peak intensities was obtained. Additionally, peaks with missing values larger than 50% were excluded. The internal standard was used for data QC (reproducibility). Metabolites were identified using the Progenesis QI automatic based on the accurate mass, MS/MS fragments, and isotope label by searching HMDB (, Lipidmaps (, and METLIN ( public databases.

Targeted metabolites analysis

Targeted metabolites analysis was conducted by an Agilent 1260 system which has been described in our former study [66]. Columns of Unitary C18 (4.6 × 250 mm, 5 μm, Acchrom) and Eclipse Plus C18 (250 mm × 4.6 mm, 5 μm, Agilent) were used. Mixed standard solutions of five anthraqunones, including emodin, aloe-emodin, rhein, chrysophanol and physcion (31.2 μgmL − 1 , 38.8 μgmL − 1 , 22 μgmL − 1 , 23.4 μgmL − 1 , and 23.4 μgmL − 1 ), and mixed standards of sennoside A and sennoside B (150 μgmL − 1 and 84mgmL − 1 ) were made. Peak area was used to calculate contents by comparing to standard solutions.

Data analysis

The data containing both negative and positive ions was analysed using the SIMCA software package (version 14.0, Umetrics, Umeå, Sweden). Principle component analysis (PCA) was used to reveal the overview classification of the samples while orthogonal projections to latent structures discriminant analysis (OPLS-DA) was used to determine the maximum separation of HA and HB samples. 7-fold cross validation and response permutation testing (RPT) were applied to avoid over-fitting. R 2 and Q 2 were calculated as 0.761 and − 0.469 after 200 times of RPT indicating that the model was accurate. In addition, univariate analysis was conducted to further confirm the significantly different metabolites. The p value and the fold change value were obtained using student’s t-test in combination with fold change analysis and the results were visualized using a volcano figure (Fig. S2). Meanwhile, the variable importance for the projection (VIP) value together with the p value were used to select the different components. The metabolites with a VIP value of > 4.0 and p < 0.05 were considered relevant for group discrimination [69].

One-way analysis of variance (ANOVA) was used with Turkey test to compare mean values for plant morphological analysis with the significant level being set as 0.05. One-way ANOVA was also used to analyze the soil properties. Targeted metabolites were compared between under canopy and open habitats according to Mann-Whitney nonparametric test.

Network study was conducted by searching the 31 most changed metabolites. Swiss ADME ( [70] was used to filter the metabolites based on their ADME (absorption, distribution, metabolism, and excretion) assessments. Metabolites with high gatrointestinal absorption index (GI absorption) and at least three ‘yes’ in five druglikeness indexes (they were Lipinski, Ghose, Veber, Egan and Muegge, respectively) were left for the target prediction. Then, Swiss Target Prediction tool was employed to predict potential gene targets of the filtered metabolites from Swiss ADME tool [71]. Targets with probability larger than 0 were used in the network contribution with their common name. At last, network files contained 6 metabolites and 208 targets were imported into the Cytoscape 3.8.2 software for network analysis. The network figure was contributed by Cytoscape 3.8.2 [72].