Orange agoseris (Agoseris aurantiaca [Hook.] Greene) belongs to the Cichorieae tribe of the Asteraceae family (Lee et al. 2003).
Asteraceae – Aster family
NRCS Plant Code
AGAU2 (USDA NRCS 2020).
The following varieties are currently recognized by the Flora of North America (Baird 2006): Agoseris aurantiaca var. aurantiaca (Hook.) Greene and A. a. var. purpurea (A. Gray).
Agoseris species are widespread with high levels of morphological and regional variability (Baird 2006). This variability makes identification difficult and many inaccurate descriptions exist in the literature. Hybridization and intermediate forms are also common within the genus. Many synonyms for Agoseris species have resulted from the naming of variant populations: Troximon aurantiacum (Baird 2006).
Variety aurantiaca: A. angustissima (Greene), A. arachnoidea Rydberg, A. carnea Rydberg, A. gaspensis Fernald, A. gracilens (A. Gray) Greene, A. g. var. greenei (A. Gray) S.F. Blake, A. greenei (A. Gray) Rydberg, A. howellii Greene, A. lackschewitzii (Douglas) M. Henderson & R.K. Moseley, A. nana Rydberg, A. naskapensis J. Roussea & Raymond, A. prionophylla Greene, A. subalpina G.N. Jones, A. vulcanica Greene (Baird 2006).
Variety purpurea: Macrorhynchus purpureus A. Gray, A. arizonica (Greene) Greene, A. attenuata Rydberg, A. aurantiaca subsp. purpurea (A. Gray) G.W. Douglas, A. confinis Greene, A. frondifera Osterhout, A. glauca (Pursh) Rafinesque var. cronquistii S.L. Welsh, A. graminifolia Greene, A. longirostris Greene, A. purpurea (A. Gray) Greene, A. purpurea var. arizonica (Greene) G.L. Wittrock, A. roseata Rydberg, A. rostrata Rydberg (Baird 2006).
Orange agoseris, mountain agoseris, mountain dandelion, orange mountain dandelion (Shaw 1995; Baird 2006; Blackwell 2006).
Chromosome numbers are: 2n = 18, 34, 36; 2n = 18 for variety aurantiaca and 2n = 18, 34, 36 for variety purpurea (Löve 1968; Baird 2006). 2n = 17 was reported for plants collected in Arizona (Keil et al. 1988).
Variety aurantiaca has formed hybrids with A. glauca, A. grandiflora, A. monticola, and A. parviflora. Variety purpurea has hybridized with A. parviflora, A. glauca, and A. g. var. dasycephala. A. × montana Osterhout is the recognized hybrid of A. a. var. purpurea and A. g. var. dasycephala that occupies high-elevation sites in the Rocky Mountains of Colorado (Baird 2006).
Orange agoseris is widespread throughout the Rocky Mountains and Intermountain West (Cronquist 1994; Welsh et al. 2016), occurring in all western states and Canadian territories west of Colorado. It is also found in South Dakota and a disjunct population occurs in Quebec (Baird 2006). Variety aurantiaca is the most common variety occurring throughout the range identified for the species. Variety purpurea is common in the Colorado Plateau and southern Rocky Mountains and occurs in Nevada, Utah, Wyoming, Colorado, Arizona, and New Mexico (Baird 2006).
Habitat And Plant Associations
Orange agoseris occupies dry to wet sites ranging from lowland grasslands and shrublands to subalpine meadows and alpine tundra (Baird 2006; Hitchcock and Cronquist 2018). It is often found along roadsides and in other disturbed sites (Baird 2006). In Utah, variety aurantiaca occurs in sagebrush (Artemisia spp.), mountain brush, pinyon-juniper (Pinus–Juniperus spp.), and alpine meadow communities, and variety purpurea occurs in mountain brush, quaking aspen (Populus tremuloides), quaking aspen-fir (Abies spp.), and spruce (Picea spp.)-fir communities (Welsh et al. 2016). Orange agoseris is often found in association with big sagebrush (Artemisia tridentata), rabbitbrush (Chrysothamnus spp., Ericameria spp.), perennial grasses, and pinyon-juniper from valleys and lowlands to as high as 10,800 ft (3,300 m) in the mountains of Utah (Jensen 2007). In California, it occurs along streams and in meadows (Fig. 1), shrublands, and moist grassland openings in montane conifer forests (Munz and Keck 1973; Hickman 1993). In the Great Basin, Balckwell (2006) notes orange agoseris occurs in seep areas in rocks, on grassy slopes, and in openings in montane communities.
The following studies describe some of the species and plant communities associated with orange agoseris at sporadic sites throughout its range. Descriptions of plant associations from orange agoeris’ southern-most range are lacking. In Alberta, orange agoseris occurs with alpine timothy (Phleum alpinum) and tufted hairgrass (Deschampsia caespitosa) in alpine meadows at the forest-tundra ecotone (Heusser 1956). Orange agoseris occurs in several subalpine meadows in the Olympic Mountains in Washington. It was most common in mesic Idaho fescue (Festuca idahoensis) grasslands occupying dry south- to west-facing slopes from 5,150 to 5,550 ft (1,570–1,690 m) elevations in the northeastern part of the mountain range. These grasslands occupy relatively steep slopes and are snow-free by mid-June. Orange agoseris was next most common in cushion plant communities on moderately steep to steep south- to southwest facing slopes between 5,050 to 5,250 ft (1,540–1,600 m). The cushion plant communities occupy windy sites that are snow-free by early May and experience frequent spring freezing and thawing (Kuramoto and Bliss 1970). In Mount Rainier National Park, Washington, orange agoseris occurs in Sunrise Meadows with poorly developed soils dominated by greenleaf fescue (F. viridula) and broadleaf lupine (Lupinus latifolius) (Frank and del Moral 1986).
Figure 1. Orange agoseris growing in a meadow in California. Photo: J. Pawek, CalPhotos, 2012.
In the Cascade Range in Oregon, orange agoseris grows in montane meadow openings in conifer forests dominated by lodgepole pine (Pinus contorta) and grand fir (Abies grandis). Dominant species in the meadows include California brome (Bromus carinatus), blue wildrye (Elymus glaucus), and wild strawberries (Fragaria spp.) (Halpern et al. 2014). On Fairview Mountain in southeastern Lane County, Oregon, orange agoseris occurs along roadsides between 5,000 and 6,000 ft (1,500–1,800 m) elevation, where the growing season is short and mountain hemlock (Tsuga mertensiana) and Alaska cedar (Chamaecyparis nootkatensis) are common (Baker 1951). In the Wallowa Mountains, orange agoseris occurs with western moonwort (Botrychium hesperium) in mesic meadows or forest edges between 5,035 and 5,450 ft (1,535–1,660 m) in steep calcareous canyons of the Lostine River drainage (Zika et al. 2002). Orange agoseris occurs infrequently along roadsides and in dry meadows in the Tenderfoot Creek Experiment Forest in the Little Belt Mountains of Montana (Mincemoyer and Birdsall 2006). In the Wind River Mountains of Wyoming, it grows in tufted hairgrass-flat-top pussytoes (Antennaria corymbosa) meadows surrounded by subalpine forests. These meadows are wet after snowmelt and dry as summer progresses (Cole and Monz 2002). In Gunnison County, Colorado, orange agoseris is a common forb in subalpine meadows at about 11,200 ft (3,400 m) elevation in the Virginia Basin (Huntly 1987). On Arizona’s Kiabab Plateau at an elevation of 8,400 ft (2,560 m), orange agoseris occurs in meadow vegetation dominated by mountain muhly (Muhlenbergia montana), squirreltail (Elymus elymoides subsp. elymoides), and pine dropseed (Blepharoneuron tricholepis) (Merkle 1962).
Orange agoseris occurs from 650 to 11,800 ft (200–3,600 m) in elevation. Variety aurantiaca occupies this entire range, while variety purpurea is absent at lower elevations, occurring at mid- and higher elevations from 5,900 to 11,800 ft (1,800–3,600 m) (Baird 2006). The elevation range for the species in the Great Basin is 6,500 to 11,500 ft (2,000–3,500 m) (Blackwell 2006) and in California it is 4,900 to 11,500 ft (1,500–3,500 m) (Hickman 1993). In Utah, variety aurantiaca’s elevation range is 4,510 to 11,240 ft (1,375–3,425 m) (Welsh et al. 2016).
The soils described in orange agoseris habitats throughout its range are highly variable in texture and depth (Fig. 2). Soil moisture requirements are also broad (Gavin and Brubaker 1999), although dry to moist, well-drained soils are generally preferred (Jensen 2007; LBJWC 2021; Plants for a Future 2020). On Arizona’s Kiabab Plateau, orange agoseris grows in meadows on shallow limestone sands (Merkle 1962). Both orange agoseris varieties occur in high-altitude meadows in Boulder Park, northern Colorado, which are concentrated on moist, gravelly soils. These high-altitude meadows are well supplied with moisture, but the climate is severe, and frosts occur nearly every week (Reed 1917). In Oregon’s Cascade Range, orange agoseris grows in meadow openings of montane conifer forests where soils are deep (>5.6 ft [1.7 m]), very fine sandy loams (Halpern et al. 2014). In the Crested Butte area of west-central Colorado (10,500–11,500 ft [3,200–3,500 m]), orange agoseris averaged 0.5% cover on 60-year-old burn scars on heavy clay-textured soils (Langenheim 1962). On Deer Ridge in Olympic National Park, Washington (5,250–5,580 ft [1,600–1,700 m]), orange agoseris occurs in meadows with weakly developed, sandy soils. Moisture content of these soils ranged from 8.5 to 25.4% between mid-June to early September (del Moral 1983a).
Figure 2. Orange agoseris growing at a rocky site in California. Photo: G.A. Monroe, CalPhotos, 1987.
Orange agoseris grows in nutrient poor soils, but its abundance may be greater and growth better on soils with higher nutrient content or when nutrients are added. Ant (Formica canadensis) mound and nonmound vegetation were compared in south-facing meadows near Gothic, Colorado (9,500 ft [2,900 m]) (Culver and Beattie 1983). Density of orange agoseris on mounds (1.1 stems) was about twice that on nonmounds (0.5 stems). Ant mounds were 4 to 5 years old, and all but one was inactive. Phosphorus, potassium, zinc, and iron contents were significantly greater on mounds than nonmounds (P < 0.05), but organic matter was significantly less on mounds than nonmounds (Culver and Beattie 1983). On Deer Ridge in Olympic National Park, Washington (5,250–5,580 ft [1,600–1,700 m]), total biomass of agoseris (orange agoseris and pale agoseris [Agoseris glauca]) increased significantly (P < 0.05) when dominant species were removed (cut at the base up to 4 times/season) and fertilizer was added (0.02 lb/ft2 [80 g/m2], 18% N, 24% P, 6% K) (del Moral 1983a). In subalpine meadows near the Rocky Mountain Biological Laboratory in western Colorado, nitrogen deposition rates are low (averaging 0.4 g nitrate, 0.06 g ammonium/yr). Treatments adding low (1 g N/m2/yr) and high (20 g N/m2/yr) levels of nitrogen did not improve per flower female reproduction of orange agoseris (Burkle and Irwin 2010).
Orange agoseris is a perennial scapose herb (Fig. 3) from a simple to branched caudex and strong, deep taproot (USFS 1937; Munz and Keck 1973; Welsh et al. 2016). Herbage is glabrous to villous and exudes a milky juice when damaged (Spellenberg 2001; Welsh et al. 2016). Leaves are entirely basal, erect to decumbent with leaf blades 1.4 to 15 in (3.5–38 cm) long and 0.1 to 1.2 in (0.3–3 cm) wide (Baird 2006). Leaf blades are narrow, linear to oblanceolate, and broadest at the middle (Spellenberg 2001). Leaf margins range from entire or with several teeth or lobes occurring in two to four pairs (Baird 2006). Flowers are produced individually at the ends of upright leafless stalks (Spellenberg 2001).
Figure 3. Orange agoseris plant in flower growing in Blaine County, Idaho. Photo: ©2017 Barry Rice.
Plants in flower can reach 32 in (80 cm) tall but are often 16 in (41 cm) tall or less (Hermann 1966). Orange agoseris produces flower heads that are about an inch (2.5 cm) wide (Fig. 4) and are comprised of 15 to 100 ray-like florets (Spellenberg 2001; Baird 2006; Welsh et al. 2016). The florets are ligulate with five lobes at the tip and uniformly fertile (Delmatier and Fertig 2014). Inner ligulate florets are shorter than the outer florets (Spellenberg 2001). The corolla is orange, occasionally yellow orange or pink, and dries to purple or burnt orange (Hickman 1993; Hitchcock and Cronquist 2018). The involucre, which encloses the flower head is 0.5 to 1.2 in (1.2–3 cm) high, 0.4 to 1.6 in (1–4 cm) wide, and often tomentose (Fig. 5). Outer, often purple-tinged bracts, can be glabrate and are shorter and wider than the inner bracts (Welsh et al. 2016; Hitchcock and Cronquist 2018). Ray-like flowers produce stamens with anthers that are sagittate at the base with rounded tips. Pistils have linear style branches.
Orange agoseris produces cylindric cypselae fruits, but these are often referred to as achenes in the literature. The cypselae have 0.4 to 0.6-in (0.9–1.5 cm) long pappi of capillary bristles arranged in a two or three series (Baird 2006; Welsh et al. 2016). Cypselae are 10-ribbed, 4 to 9 mm long, with slender beaks. The beaks are often more than half as long as the cypselae bodies (Munz and Keck 1973; Welsh et al. 2016; Hitchcock and Cronquist 2018).
Figure 4. Orange agoseris flower head with lobed ligulate florets the inner of which being shorter than the outer. Photo: ©Al Schneider, swcoloradowildflowers.com.
Figure 5. Tomentose orange agoseris involucre enclosing the flower head. Photo: C. Wagner, CalPhotos, 2014.
Varieties. The Flora of North America suggests that varieties are best distinguished by the involucre bracts and the beaks and ribs of cypselae (Baird 2006).
Variety aurantiaca produces leaves that often have entire margins (Baird 2006; Welsh et al. 2016). Flower stems are often shorter than the leaves (Baird 2006). Involucre bracts can be more than 0.8 in (2 cm) long and are subequal, slender and tapering (Hermann 1966; Welsh et al. 2016). Cypselae have abruptly tapered beak ends and ribs that are often thicker distally (Baird 2006). Variety aurantiaca is often much taller than variety purpurea (Welsh et al. 2016).
Variety purpurea produces leaves with typically dentate margins (Baird 2006; Welsh et al. 2016). Flower stems are often shorter than the leaves (Baird 2006). Involucre bracts are overlapping, broad, with blunt rounded tips (Hermann 1966; Welsh et al. 2016). Cypselae have gradually tapered beak ends and ribs that are weakly and uniformly ridged but not thickened distally (Baird 2006).
Orange agoseris flowers from May to September (Baird 2006; Jensen 2007), and seed typically ripens from May to September (Jensen 2007; PFAF 2021). Plants produce one to three seed heads each year but may produce more flowering stalks if soil moisture is not limited (Fig. 6) (Jensen 2007).
Figure 6. Single orange agoseris plant producing both flowers and seeds (indeterminate growth). Photo: ©Al Schneider, swcoloradowildflowers.com.
Flowers have both stamens and pistils and are pollinated by insects (PFAF 2021).
Pollinators from the following families: Anthomyiidae, Halictidae, Hesperiidae, Lycaenidae, Muscidae, Nymphalidae, Pieridae, Sarcophagidae, Syrphidae, and Tachinidae interacted with orange agoseris flowers in high-elevation meadows (9, 500 ft [2,900 m]) near Crested Butte, Colorado. Flower-pollinator interactions were observed for 3 years (June through August 2005–2007) during good weather from 9 am to 4 pm (Burkle and Irwin 2009). Orange agoseris was an important nectar source for 27 butterfly species in the Crested Butte area of Colorado, which was monitored for four flight seasons (Sharp et al. 1974).
A strong taproot and wind-dispersed seed (Fig. 6) suggests orange agoseris is disturbance adapted (Shaw 1995). Plants are considered shade intolerant (PFAF 2021) and early colonizers of disturbed sites (Titus et al. 1998; Brown et al. 2003). The life span of orange agoseris has been described as short- (Eldredge et al. 2013) to long-lived (Romme et al. 2016).
Several studies reported orange agoseris appeared in early primary succession of severely disturbed sites (Frank and del Moral 1986; Titus et al. 1998; Brown et al. 2003). It colonized a denuded subalpine meadow on poorly developed soils at Mount Rainier National Park, Washington. Denuded sites resulted from cabin removal which was followed by bulldozing. Orange agoseris cover was 0.1% on bulldozed sites within 10 years, which was just slightly less than its cover in nearby undisturbed meadows (Frank and del Moral 1986). Orange agoseris colonized an abandoned mine site in southern Montana (Brown et al. 2003). The mine, operated for copper, gold, and silver extraction, was abandoned in 1952. By 1972, plant colonization was limited. Following treatments of lime, fertilizer, and a mix of native and nonnative grasses in 1976, orange agoseris appeared on treated but not untreated mine spoils (Brown et al. 2003). Following the 1980 eruption of Mt. St. Helens in Washington, orange agoseris was found soon after the eruption even on sites with thick mud and little vegetation recovery or colonization (del Moral 1983b). In later sampling, 13 to 14 years after the eruption, orange agoseris was infrequently found at sites where vegetation was essentially eliminated. It was common at refugia sites where some vegetation survived (Titus et al. 1998).
Disturbed sites are common habitats for orange agoseris, and its abundance is often greater on harvested, grazed, or burned sites than on undisturbed sites. In a subalpine huckleberry (Vaccinium spp.) field at 1,200 feet (4,000 ft) elevation on Mount Adams in southwestern Washington, effects of heavy sheep grazing, cutting and burning, and fire alone were evaluated. Cover of orange agoseris was greater in treated (0.2–0.4%) than untreated (0–0.1%) sites in all 4 years of post-treatment monitoring (Minore et al. 1979). In the Snowy Range of southeastern Wyoming, orange agoseris occurred on 30- to 50-year-old clearcuts (0.01–0.09% average cover) but was absent from mature lodgepole pine (Pinus contorta) montane and subalpine forests (subalpine fir [Abies lasiocarpa] and Engelmann spruce [Picea engelmanii]). The basal area of trees was 35 to 61 ft2/acre (8–14 m2/ha) in harvested and 17,091 to 27,825 ft2/acre (3,925–6,390 m2/ha) in mature forests (Selmants and Knight 2003).
In grasslands surrounded by ponderosa pine (Pinus ponderosa) forests on Arizona’s Mogollon Rim (~7,200 ft [2,200 m] elevation), the relative abundance of orange agoseris varied between grazed and ungrazed sites protected from cattle, deer (Odocoileus spp.), and elk (Cervus canadensis) for 8 to 9 years. Relative abundance of orange agoseris was 0.91 on grazed and 0.14 on protected plots at site one, 0.14 on grazed and 1.1 on protected plots at site two, and 0 on grazed and 2.16 on protected plots at site three. Cattle use resulted in about 70% biomass consumption (Rambo and Faeth 1999).
In a montane meadow in Oregon’s Cascade Range, the frequency and cover of orange agoseris were similar at the interior of pile burn and unburned plots 7 years after treatments. Orange agoseris frequency was 3.3% and cover 0.1% at the center of pile burn scars and frequency was 3.3% and cover 0.3% in unburned vegetation adjacent to the burn scar (Halpern et al. 2014). Following the 1988 fires in Yellowstone National Park, orange agoseris was more common on recently burned sites. It was present on 25% of burned plots in 1999 and just 1% of burned plots in 2012 (Romme et al. 2016). In krummholz communities on Sourdough Ridge in Washington’s North Cascades National Park, cover and frequency of orange agoseris were greater on 29-year-old burned (1% cover, 4–10% frequency) than unburned sites (0%-trace cover, 0–5% frequency) (Douglas and Ballard 1971).
Wildlife And Livestock Use
Orange agoseris is eaten by a variety of livestock and wildlife species, and high levels of use were reported for domestic sheep and northern pocket gophers (Thomomys talpoides). Hermann (1966) indicated that orange agoseris was fair to good livestock forage, and variety purpurea was highly palatable to sheep and cattle on the Manti-La Sal National Forest. In feeding observations on twoneedle pinyon-Utah juniper (Pinus edulis–Juniperus osteosperma) winter range in the Piceance Basin in Colorado, orange agoseris made up to 2% of the diets of tame mule deer (Odocoileus hemionus) from October through April (Bartmann 1983).
Small mammal use of orange agoseris was reported in Utah and Colorado. On Utah’s Wasatch Plateau, removal of orange agoseris was observed on five northern pocket gopher mounds (Aldous 1951). On the Black Mesa of west-central Colorado, orange agoseris was among the 10 most important forbs in diets of northern pocket gophers trapped in Idaho fescue-dominated grasslands (Carleton 1966). From stomach content analyses of 397 individuals, researchers found that northern pocket gophers fed on leaves, stems, and roots. Based on 2 years of monthly use from June through October, use was greatest in October when stems and leaves made up 6.4% and roots made up 17.7% of diets by volume (Ward and Keith 1962). In the mountains of west-central Colorado where golden-mantled ground squirrels (Citellus lateralis lateralis) and least chipmunks (Neotamias minimus) are sympatric, orange agoseris was ranked as edible on a scale from highly edible, edible, sometimes edible, and never edible (Carleton 1966).
Agoseris species are eaten by and can be important in the diets of greater sage-grouse (Centrocercus urophasianus) (Martin et al. 1951; Pennington et al. 2016). In a review of the use of forbs by greater sage-grouse, Agoseris species were among the most important for females in spring and for adults and juveniles in summer in sagebrush habitats in Oregon, Idaho, and Nevada (Pennington et al. 2016).
Orange agoseris was an important nectar source for 27 butterfly species in the Crested Butte area of Colorado, which were monitored for four flight seasons (Sharp et al. 1974).
Orange agoseris was and is used as a food and medicine by Indigenous peoples. It is a potherb used to make teas and wines (Shaw 1995). The Karuk tribe of California sucked the juice out of the plant near the crown and chewed it like gum (Schenck and Gifford 1952). Gosiute Indians of Utah and Nevada ate the leaves (Chamberlin 1911). The Ramah Navajo used orange agoseris as a ceremonial medicine. An infusion of leaves or roots was taken internally or used as a lotion to treat arrow or bullet wounds, and wet leaves were rubbed on swollen arms, wrists, and ankles. Orange agoseris was considered a life medicine and provided protection from witches by the Ramah Navajo (Moerman 2003).
Although orange agoseris is not readily available as a horticultural species, it does have potential for use in low-maintenance landscaping along roadsides, at rest areas and campgrounds. It produces attractive orange flowers and spreads by wind-dispersed seeds.
Orange agoseris has many traits that make it a good species for restoring native habitats. It is a colonizer of early successional and disturbed habitats. It is attractive to pollinators (Eldredge et al. 2013). The species has broad moisture tolerance (Gavin and Brubaker 1999) and a wide-ranging elevation tolerance (Baird 2006) suggesting it may be adaptable to changing climates or transitional sites.
Developing A Seed Supply
For restoration to be successful, the right seed needs to be planted in the right place at the right time. Coordinated planning and cooperation is required among all partners first to select appropriate species and seed sources; these are determined by current or projected revegetation requirements and goals (PCA 2015).
Developing a seed supply begins with seed collection from native stands. Collected seed is distributed for restoration or first increased in seed fields to provide a larger supply of seed (PCA 2015). Collection sites are determined by current or projected revegetation requirements and goals as well as through consultation with land managers and stakeholders. Production of nursery stock requires less seed than large-scale seeding operations, which may require establishment of agricultural seed production fields. Regardless of the size and complexity of any revegetation effort, seed certification is essential for tracking seed origin from collection through use (UCIA 2015).
Because empirical seed zones are not currently available for orange agoseris, generalized provisional seed zones developed by Bower et al. (2014), may be used to select and deploy seed sources. These provisional seed zones identify areas of climatic similarity with comparable winter minimum temperature and aridity (annual heat:moisture index). In Figure 7, Omernik Level III Ecoregions (Omernik 1987) overlay the provisional seeds zones to identify climatically similar but ecologically different areas. For site-specific disturbance regimes and restoration objectives, seed collection locations within a seed zone and ecoregion may be further limited by elevation, soil type, or other factors.
The Western Wildland Environmental Threat Assessment Center’s (USFS WWETAC 2017) Threat and Resource Mapping (TRM) Seed Zone application provides links to interactive mapping features useful for seed collection and deployment planning. The Climate Smart Restoration Tool (Richardson et al. 2020) can also guide revegetation planning, seed collection, and seed deployment, particularly when addressing climate change considerations.
Figure 7. Distribution of orange agoseris (black circles) based on geo-referenced herbarium specimens and observational data from 1876-2018 (CPNWH 2017; SEINet 2017; USDI USGS 2017). Generalized provisional seed zones (colored regions) (Bower et al. 2014) are overlain by Omernik Level III Ecoregions (black outlines) (Omernik 1987; USDI EPA 2018). Interactive maps, legends, and a mobile app are available (USFS WWETAC 2017; www.fs.fed.us/wwetac/threat-map/TRMSeedZoneMapper2.php?). Map prepared by M. Fisk, USDI USGS.
As of 2021, there were no orange agoseris germplasm releases.
Wildland Seed Collection
Orange agoseris plants are commonly found at low densities, so wildland seed is typically collected by hand (Jensen 2007).
Wildland Seed Certification
Verification of species and tracking of geographic source is necessary whether wildland seed is collected for immediate project use or as stock seed for cultivated increase. This official Source Identification process can be accomplished by following procedures established by the Association of Official Seed Certifying Agencies (AOSCA) Pre-Variety Germplasm Program (UCIA 2015; Young et al. 2020). Wildland seed collectors should become acquainted with state certification agency procedures, regulations, and deadlines in the states where they collect.
If wildland-collected seed is to be sold for direct use in ecological restoration projects, collectors must apply for Source-Identified certification prior to making collections. Pre-collection applications, site inspections, and species and seed amount verification are handled by the AOSCA member state agency where seed collections will be made (see listings at AOSCA.org).
If wildland seed collected by a grower or private collector is to be used as stock seed for planting cultivated seed fields or for nursery propagation (See Agricultural Seed Certification section), detailed information regarding collection site and collecting procedures must be provided when applying for certification. Photos and herbarium specimens may be required. Germplasm accessions acquired within established protocols of recognized public agencies, however, are normally eligible to enter the certification process as stock seed without routine certification agency site inspections. For contract grow-outs, however, this collection site information must be provided to the grower to enable certification.
On individual plants, seed ripens indeterminately, but across a population, seed ripens quite synchronously (Fig. 8). The first seed heads ripen as a group as do subsequent seed heads across a population. Ripening continues as moisture permits (Jensen 2007).
Seed is wind-dispersed, making the timing of collections critical. Seeds are mature when the pappus begins to spread open (Fig. 9) (Jensen 2007). The Bureau of Land Management’s Seeds of Success collection crews made 11 harvests, one from Nevada, two from Oregon, and eight from Utah from 2008 to 2010. Of the ten harvests made in 2010, the earliest was made on June 14 in Wasatch County, Utah, at an elevation of 5,485 ft (1,672 m). The latest was made on August 8 from Wasco County, Oregon, at an elevation of 4,470 ft (1,362 m). Multiple harvests were often made at the same site. The widest difference in collection dates at a single site was 16 days (June 21-July 6) (SOS BLM 2017). Jensen reported that orange agoseris seeds were typically mature and ready for harvest in May and June at elevations between 5,200 and 6,100 ft (1,600-1,900 m) in the Great Basin (Jensen 2007).
Figure 8. Orange agoseris flowering and just starting to produce seed. Photo: S. Matson, CalPhotos, 2017.
Figure 9. Mature orange agoseris seed heads dispersing seed. Photo: J. Pawek, CalPhotos, 2010.
Because orange agoseris plants typically occur in low densities, wildland seed is hand collected. Seeds are hand stripped by clasping the base of the seed head between two fingers and pulling upward while keeping the hand closed to reduce losing mature seeds (Jensen 2007).
Several collection guidelines and methods should be followed to maximize the genetic diversity of wildland collections: 1) collect seed from a minimum of 50 randomly selected plants; 2) collect from widely separated individuals throughout a population without favoring the most robust or avoiding small stature plants; and 3) collect from all microsites including habitat edges (Basey et al. 2015). General collecting recommendations and guidelines are provided in online manuals (e.g., ENSCONET 2009; USDI BLM SOS 2021).
It is critical that wildland seed collection does not impact the sustainability of native plant populations. Collectors should take no more than 20% of the viable seed available at the time of harvest (USDI BLM SOS 2021). Additionally, care must be taken to avoid the inadvertent collection of weedy species, particularly those that produce seeds similar in shape and size to those of orange agoseris.
Jensen (2007) recommends storing seed (Fig. 10) in breathable bags in cool, dry, rodent-free conditions until it can be cleaned.
Figure 10. Individual orange agoseris seeds with visible ribs and stigma and style still attached. Photo: J. Pawek, CalPhotos, 2012.
Small hand-harvested seed collections can be cleaned of larger chaff by hand sifting or screening. Seed can then be carefully rolled between two boards covered with soft leather until the pappus is removed from the seed. The final cleaning can be done using air columns or screen separators (Jensen 2007). Although seed will germinate with the stigma and style attached, removal of the pappus and other reproductive structures is necessary if orange agoseris will be used in a seed mix or be seeded using equipment (S. Jensen, USFS, personal communication, March 2021).
Orange agoseris produces orthodox seed. Seed viability was 79% for dry seed stored for 34 days at 15% relative humidity and -4 °F (-20 °C) (RBG Kew 2021, original seed viability unknown).
There is no Association of Official Seed Analysts (AOSA) rule for testing germination or AOSA protocol for examining the viability of orange agoseris seed (AOSA 2010; 2016). Tetrazolium chloride testing (TZ) should follow that suggested for other Asteraceae species, which includes cutting seeds longitudinally leaving the distal end intact, soaking for 6 to 12 hours in 0.1% TZ concentration at (86–95 °F [30–35 °C]). The embryos of viable seed will be entirely stained (AOSA 2010).
Seed germinates soon following stratification. Jensen (2007) found that following 3 weeks of stratification most seed germinated within 2 weeks of being transferred to a greenhouse. Seed was planted in trays filled with moist soil that were wrapped in plastic and placed in a cold room. Trays were then placed in a greenhouse at 70 °F (21 °C) during the day and 50 °F (10 °C) at night. Seed germinated within 5 weeks but most germinated by week 2. The optimal duration of stratification likely varies by seed source (Jensen 2007).
Germination trials conducted by the Royal Botanic Gardens, Kew (RBG Kew 2021) reported 50% germination after 70 days for unstratified seed germinated on 1% agar and exposed to an 8 hr/16 hr light/dark cycle at 41 °F (5 °C). Germination was 58% for unstratified seed germinated on 1% agar for 140 days on a 77/50 °F (25/10 °C) (8 hr light/16 hr dark).
Wildland Seed Yield And Quality
Post-cleaning seed yield and quality of seed lots collected in the Intermountain region are provided in Table 1 (USFS BSE 2017). Orange agoseris seed can generally be cleaned to high levels of purity. For fresh seed, fill is generally high, but viability is variable. Other sources report values within the range reported in Table 1 (281,215–304,772 seeds/lb [619,963–671,896 seeds/kg) (USFS GBNPP 2014; RBGKew 2021).
Table 1. Seed yield and quality of orange agoseris seed lots collected in the Intermountain region, cleaned by the Bend Seed Extractory, and tested by the Oregon State Seed Laboratory or the USFS National Seed Laboratory (USFS BSE 2017).
Seed lot characteristic
Bulk weight (lbs)
Clean weight (lbs)
Pure live seeds/lb
1 100 seed X-ray test
2 Tetrazolium chloride test
Acceptable seed purity, viability, and germination specifications vary with revegetation plans. Purity needs are highest for precision seeding equipment used in nurseries, while some rangeland seeding equipment handles less clean seed quite well.
Agricultural Seed Production
Field production of orange agoseris seed was not reported in the literature as of 2021. The indeterminate seed set, and readily wind-dispersed seed will make field production challenging (see section). Jensen and Jones (2008) attempted greenhouse seed production, but few plants flowered the first growing season and plants were not grown through their second year.
Agricultural Seed Certification
In order to minimize genetic changes in specific accessions of native species when increased in cultivated fields, it is essential to track the geographic source and prevent inadvertent hybridization or selection pressure. This is accomplished by following third party seed certification protocols for Pre-Variety Germplasm (PVG) as established by the Association of Official Seed Certification Agencies (AOSCA). AOSCA members in the U.S., Canada, and other countries administer PVG requirements and standards that track the source and generation of planting stock. Field and cleaning facility inspections then monitor stand establishment, proper isolation distances, control of prohibited weeds, seed harvesting, cleaning, sampling, testing, and labeling for commercial sales (UCIA 2015; Young et al. 2020).
Seed growers apply for certification of their production fields prior to planting and plant only certified stock seed of an allowed generation (usually less than four). The systematic and sequential tracking through the certification process requires preplanning, knowing state regulations and deadlines, and is most smoothly navigated by working closely with state certification agency personnel. See the Wildland Seed Certification section for more information on stock seed sourcing.
High orange agoseris seed yields will require pollinators (Jensen and Jones 2008), and farming operations that encourage visitation from native pollinators will likely help with seed production (Cane 2008).
The following fungi have been collected from orange agoseris: Entyloma polysporum, Leptosphaeria erigerontis, Mycosphaerella tassiana, Ovularia compacta, Phyllosticta spp., Puccinia agoseridis, P. dioicae, P. hieracii, P. troximontis, Ramularia compacta, and Sphaerotheca macularis. It is unknown if any of these fungi would be problematic in nursery or field production of orange agoseris (Farr and Rossman 2017).
Jensen (2007) produced orange agoseris plugs in the greenhouse. Seed was collected from Juab County, Utah (5,950 ft [1,800 m]). It was planted in 5.5 in3 (90 cm3) root trainers filled with two parts sieved peat, two parts vermiculite, 1 part montmorillonite clay, 1 part #20 quartz silica sand, 1 part native soil, and fertilizer with micronutrients. The potting mix was steam aerated at 140 °F (60 °C) for 30 minutes before seeds were planted vertically to a depth equal to the seed body. Seeded trays were moistened, wrapped in plastic bags, and placed in a cold room. After 3 weeks of stratification, trays were moved to a greenhouse (December), and watered as needed through establishment. The greenhouse was kept at 70 °F (21 °C) during the day and 50 °F (10 °C) at night. All germination occurred within 5 weeks, but most germination occurred in the second week. Seedlings were watered every 2 to 3 days through the active growth phase, which took at least 8 weeks (Jensen 2007).
Wildland Seeding And Planting
Reported use of orange agoseris in revegetation efforts has been limited. It was experimentally seeded in barren subalpine habitat created by the eruption of Mount St. Helens in Washington. It was one species in a mixture of 22 species seeded in October. Initial emergence was about 15%, but by September of the second post-seeding year, survival was around 1%. The seed used in this revegetation experiment was collected from nearby plants (Wood and del Moral 1987).
Funding for Western Forbs: Biology, Ecology, and Use in Restoration was provided by the USDI BLM Great Basin Native Plant Materials Ecoregional Program through the Great Basin Fire Science Exchange. Great thanks to the chapter reviewers: Scott Jensen, Botanist, US Forest Service, Rocky Mountain Research Station and Rose Lehman, Botanist, US Forest Service.
This research was supported in part by the USDA Forest Service, Rocky Mountain Research Station. The findings and conclusions in this publication are those of the authors and should not be construed to represent any official USDA or U.S. Government determination or policy.
Aldous, C.M. 1951. The feeding habits of pocket gophers (Thomomys talpoides Moorei) in the high mountain ranges of central Utah. Journal of Mammalogy. 32(1): 84-87.
Association of Official Seed Analysts [AOSA]. 2010. AOSA/SCST Tetrazolium testing handbook. Contribution No. 29. Lincoln, NE: Association of Official Seed Analysts.
Association of Official Seed Analysts [AOSA]. 2016. AOSA rules for testing seeds. Vol. 1. Principles and procedures. Washington, DC: Association of Official Seed Analysts.
Baird, G.I. 2006. Agoseris. In: Flora of North America Editorial Committee, ed. Flora of North America North of Mexico. Volume 19 Magnoliophyta: Asteridae (in part): Asteraceae, part 1. New York, NY: Oxford University Press: 323-334.
Baker, W.H. 1951. Plants of Fairview Mountain, Calapooya Range, Oregon. The American Midland Naturalist. 46(1): 132-173.
Bartmann, R.M. 1983. Composition and quality of mule deer diets on pinyon-juniper winter range, Colorado. Journal of Range Management. 36(4): 534-541.
Basey, A.C.; Fant, J.B.; Kramer, A.T. 2015. Producing native plant materials for restoration: 10 rules to collect and maintain genetic diversity. Native Plants Journal. 16(1): 37-53.
Blackwell, L.R. 2006. Great Basin wildflowers: A guide to common wildflowers of the high deserts of Nevada, Utah, and Oregon. Helena, MT: Morris Book Publishing. 288 p.
Bower, A.D.; St. Clair, J.B.; Erickson, V. 2014. Generalized provisional seed zones for native plants. Ecological Applications. 24(5): 913-919.
Brown, R.W.; Amacher, M.C.; Mueggler, W.F.; Kotuby-Amacher, J. 2003. Reestablishing natural succession on acidic mine spoils at high elevation: Long-term ecological restoration. Res. Pap. RMRS-RP-41. U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station. 49 p.
Burkle, L.; Irwin, R. 2009. The importance of interannual variation and bottom-up nitrogen enrichment for plant-pollinator networks. Oikos. 118(12): 1816-1829.
Burkle, L.A.; Irwin, R.E. 2010. Beyond biomass: Measuring the effects of community-level nitrogen enrichment on floral traits, pollinator visitation and plant reproduction. Journal of Ecology. 98(3): 705-717.
Cane, J.H. 2008. Pollinating bees crucial to farming wildflower seed for U.S. habitat restoration. In: James, R.; Pitts-Singer, T., eds. Bees in agricultural ecosystems. Oxford, UK: Oxford University Press: 48-64.
Carleton, W.M. 1966. Food habits of two sympatric Colorado sciurids. Journal of Mammalogy. 47(1): 91-103.
Chamberlin, R.V. 1911. The ethno-botany of the Gosiute Indians. American Anthropological Association. 63(1): 329-405.
Cole, D.N.; Monz, C.A. 2002. Trampling disturbance of high-elevation vegetation, Wind River Mountains, Wyoming, U.S.A. Arctic, Antarctic, and Alpine Research. 34(4): 365-376.
Consortium of Pacific Northwest Herbaria [CPNWH]. 2017. Agoseris aurantiaca. Seattle, WA: University of Washington Herbarium, Burke Museum of Natural History and Culture. http://www.pnwherbaria.org/ [Accessed 2020 October 25].
Cronquist, A. 1994. Volume five: Asterales. In: Cronquist, A.; Holmgren, A.H.; Holmgren, N.H.; Reveal, J.L.; Holmgren, P.K., eds. Intermountain flora: Vascular plants of the Intermountain West, U.S.A. Bronx, NY: The New York Botanic Garden. 496 p.
Culver, D.C.; Beattie, A.J. 1983. Effects of ant mounds on soil chemistry and vegetation patterns in a Colorado montane meadow. Ecology. 64(3): 485-492.
del Moral, R. 1983a. Competition as a control mechanism in subalpine meadows. American Journal of Botany. 70(2): 232–245.
del Moral, R. 1983b. Initial recovery of subalpine vegetation on Mount St. Helens, Washington. The American Midland Naturalist. 109(1): 72–80.
Delmatier, C.; Fertig, W. 2014. Plant of the week: Orange mountain dandelion (Agoseris aurantiaca). U.S. Department of Agriculture, U.S. Forest Service. https://www.fs.fed.us/wildflowers/plant-of-the-week/agoseris_aurantiaca.shtml [Accessed 2021 February 3].
Douglas, G.W.; Ballard, T.M. 1971. Effects of fire on alpine plant communities in the North Cascades, Washington. Ecology. 52(6): 1058-1064.
Eldredge, E.; Novak-Echenique, P.; Heater, T.; Mulder, A.; Jasmine, J. 2013. Plants for pollinator habitat in Nevada. Tech. Note NV 57. Reno, NV: U.S. Department of Agriculture, Natural Resources Conservation Service. 65 p.
European Native Seed Conservation Network [ENSCONET]. 2009. ENSCONET seed collecting manual for wild species. Edition 1: 32 p.
Farr, D.F.; Rossman, A.Y. 2017. Fungal databases, U.S. National Fungus Collections. U.S. Department of Agriculture, Agricultural Research Service. https://nt.ars-grin.gov/fungaldatabases/.
Frank, D.A.; del Moral, R. 1986. Thirty-five years of secondary succession in a Festuca viridula–Lupinus latifolius dominated meadow at Sunrise, Mount Rainier National Park, Washington. Canadian Journal of Forest Research. 64(6): 1232-1236.
Gavin, D.G.; Brubaker, L.B. 1999. A 6000-year soil pollen record of subalpine meadow vegetation in the Olympic Mountains, Washington, USA. Journal of Ecology. 87(1): 106-122.
Halpern, C.B.; Antos, J.A.; Beckman, L.M. 2014. Vegetation recovery in slash-pile scars following conifer removal in a grassland-restoration experiment. Restoration Ecology. 22(6): 731-740.
Hermann, F. 1966. Notes on western range forbs: Cruciferae through Compositae. Agric. Handb. 293. Washington, DC: U.S. Department of Agriculture, Forest Service. 365 p.
Heusser, C.J. 1956. Postglacial environments in the Canadian Rocky Mountains. Ecological Monographs. 26(4): 263-302.
Hickman, J.C., ed. 1993. The Jepson manual: Higher plants of California. Berkeley, CA: University of California Press. 1400 p.
Hitchcock, C.L.; Cronquist, A. 2018. Flora of the Pacific Northwest: An illustrated manual. 2nd Ed. Giblin, D.E.; Legler, B.S.; Zika, P.F.; Olmstead, R.G., eds. Seattle, WA: University of Washington Press. 882 p.
Huntly, N.J. 1987. Influence of refuging consumers (Pikas: Ochotona princeps) on subalpine meadow vegetation. Ecology. 68(2): 274-283.
Jensen, S.; Jones, C. 2008. Plant materials and development, cultural practices for Agoseris and Lupinus. In: Shaw, N.; Pellant, M., eds. Great Basin Native Plant Selection and Increase 2007 Progress Report. U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 4-6.
Jensen, S.L. 2007. Propagation protocol for production of container (plugs) Agoseris aurantiaca (Hook.) Greene, plants 5.5 cu. in. root trainers. U.S. Department of Agriculture, Forest Service, Shrub Sciences Laboratory Provo, Utah. National Center for Reforestation, Nurseries, and Genetic Resources. http://npn.rngr.net/propagation/protocols [Accessed 2020 November 20].
Keil, D.J.; Luckow, M.A.; Pinkava, D.J. 1988. Chromosome studies in Asteraceae from the United States, Mexico, the West Indies, and South America. American Journal of Botany. 75(5): 652-668.
Kuramoto, R.T.; Bliss, L.C. 1970. Ecology of subalpine meadows in the Olympic Mountains, Washington. Ecological Monographs. 40(3): 317-347.
Lady Bird Johnson Wildflower Center [LBJWC]. 2021. Agoseris aurantiaca (Hook.) Greene. Native Plant Database. Austin, TX: Lady Bird Johnson Wildflower Center. https://www.wildflower.org/plants-main [Accessed 2021 January 12].
Langenheim, J.H. 1962. Vegetation and environmental patterns in the Crested Butte Area, Gunnison County, Colorado. Ecological Monographs. 32(3): 249-285.
Lee, J., Baldwin, B.G.; Gottlieb, L.D. 2003. Phylogenetic relationships among the primarily North American genera of Cichorieae (Compositae) based on analysis of 18s-26s nuclear rDNA its and ETS sequences. Systematic Botany. 28(3): 616-626.
Löve, Á. 1968. Iopb chromosome number reports. XVII. Taxon.
Martin, A.C.; Zim, H.S.; Nelson, A.L. 1951. American wildlife and plants: A guide to wildlife food habits. New York, NY: Dover Publications. 500 p.
Merkle, J. 1962. Plant communities of the Grand Canyon area, Arizona. Ecology. 43(4): 698-711.
Mincemoyer, S.A.; Birdsall, J.L. 2006. Vascular flora of the Tenderfoot Creek Experimental Forest, Little Belt Mountains, Montana. Madrono. 53(3): 211-222.
Minore, D.; Smart, A.W.; Dubrasich, M.E. 1979. Huckleberry ecology and management research in the Pacific Northwest. Gen. Tech. Rep. PNW-93. U.S. Department of Agriculture, Forest Service, Pacific Northwest Forest and Range Experiment Station. 50 p.
Moerman, D. 2003. Native American ethnobotany: A database of foods, drugs, dyes, and fibers of Native American peoples, derived from plants. Dearborn, MI: University of Michigan. http://naeb.brit.org/.
Munz, P.A.; Keck, D.D. 1973. A California flora and supplement. Berkeley, CA: University of California Press. 1905 p.
Omernik, J.M. 1987. Ecoregions of the conterminous United States. Map (scale 1:7,500,000). Annals of the Association of American Geographers. 77(1): 118-125.
Pennington, V.E.; Schlaepfer, D.R.; Beck, J.L.; Bradford, J.B.; Palmquist, K.A.; Lauenroth, W.K. 2016. Sagebrush, greater sage-grouse, and the occurrence and importance of forbs. Western North American Naturalist. 76(3): 298-312.
Plant Conservation Alliance [PCA]. 2015. National seed strategy for rehabilitation and restoration 2015-2020. Washington, DC: U.S. Department of the Interior, Bureau of Land Management. 52 p.
Plants for a Future [PFAF]. 2021. https://pfaf.org/user/Default.aspx
Rambo, J.L.; Faeth, S.H. 1999. Effect of vertebrate grazing on plant and insect community structure. Conservation Biology. 13(5): 1047-1054.
Reed, E.L. 1917. Meadow vegetation in the montane region of northern Colorado. Bulletin of the Torrey Botanical Club. 44(2): 97-109.
Richardson, B.; Kilkenny, F.; St. Clair, B.; Stevenson-Molnar, N. 2020. Climate Smart Restoration Tool. https://climaterestorationtool.org/csrt/.
Romme, W.H.; Whitby, T.G.; Tinker, D.B.; Turner, M.G. 2016. Deterministic and stochastic processes lead to divergence in plant communities 25 years after the 1988 Yellowstone fires. Ecological Monographs. 86(3): 327-351.
Royal Botanic Gardens, Kew [RBG Kew]. 2020. Seed Information Database (SID). Version 7.1. http://data.kew.org/sid/ [Accessed 2020 February 16].
Schenck, S.M.; Gifford, E.W. 1952. Karok ethnobotany. Anthropological Records. 13(6): 377-392.
SEINet – Regional Networks of North American Herbaria Steering Committee [SEINet]. 2017. SEINet Regional Networks of North American Herbaria. Symbiota.org/docs/seinet.
Selmants, P.C.; Knight, D.H. 2003. Understory plant species composition 30-50 years after clearcutting in southeastern Wyoming coniferous forests. Forest Ecology and Management. 185(3): 275-289.
Sharp, M.A.; Parks, D.R.; Ehrlich, P.R. 1974. Plant resources and butterfly habitat selection. Ecology. 55(4): 870-875.
Shaw, R.J. 1995. Utah wildflowers: A field guide to northern and central mountains and valleys. Logan, UT: Utah State University Press. 218 p.
Spellenberg, R. 2001. National Audubon Society field guide to North American wildflowers: Western region, revised edition. New York, NY: Alfred A. Knopf, Inc. 862 p.
Titus, J.H.; Moore, S.; Arnot, M.; Titus, P.J. 1998. Inventory of the vascular flora of the blast zone, Mount St. Helens, Washington. Madroño. 45(2): 146-161.
U.S. Department of Agriculture, Forest Service [USFS]. 1937. Range Plant Handbook. Washington, DC: U.S. Department of Agriculture, Forest Service. 816 p.
U.S. Department of Agriculture, Forest Service, Great Basin Native Plant Project [USFS GBNPP]. 2014. Seed weight table calculations made in-house. Report on file. Boise, ID: U.S. Department of Agriculture, Forest Service, Boise Aquatic Sciences Laboratory. Available: https://www.fs.fed.us/rm/boise/research/shrub/Links/Seedweights.pdf
USDA Forest Service, Bend Seed Extractory [USDA FS BSE]. 2017. Nursery Management Information System Version 4.1.11. Local Source Report 34-Source Received. Bend, OR: U.S. Department of Agriculture, Forest Service, Bend Seed Extractory.
USDA Forest Service, Western Wildland Environmental Threat Assessment Center [USFS WWETAC]. 2017. TRM Seed Zone Applications. Prineville, OR: U.S. Department of Agriculture, Forest Service, Western Wildland Environmental Threat Assessment Center. https://www.fs.fed.us/wwetac/threat-map/TRMSeedZoneMapper.php.
USDA Natural Resources Conservation Service [USDA NRCS]. 2020. The PLANTS Database. Greensboro, NC: U.S. Department of Agriculture, Natural Resources Conservation Service, National Plant Data Team. https://plants.usda.gov/java.
USDI Bureau of Land Management, Seeds of Success [USDI BLM SOS]. 2016. Bureau of Land Management technical protocol for the collection, study, and conservation of seeds from native plant species for Seeds of Success. Washington, DC: USDI Bureau of Land Management. 37 p.
USDI Bureau of Land Management, Seeds of Success [USDI BLM SOS]. 2017. Seeds of Success collection data. Washington, DC: U.S. Department of the Interior, Bureau of Land Management, Plant Conservation Program.
USDI Environmental Protection Agency [USDI EPA]. 2018. Ecoregions. Washington, DC: U.S. Environmental Protection Agency. https://www.epa.gov/eco-research/ecoregions.
USDI Geological Survey [USDI USGS]. 2017. Biodiversity Information Serving Our Nation (BISON). U.S. Geological Survey. https://bison.usgs.gov/#home.
Utah Crop Improvement Association [UCIA]. 2015. How to be a seed connoisseur. Logan, UT: UCIA, Utah Department of Agriculture and Food, Utah State University and Utah State Seed Laboratory. 16 p.
Ward, A.L., Keith, J.O. 1962. Feeding habits of pocket gophers on mountain grasslands, Black Mesa, Colorado. Ecology. 43(4): 744-749.
Weber, W.A. 1976. Rocky Mountain flora. Boulder, CO: Colorado Associated University Press. 479 p.
Welsh, S.L.; Atwood, N.D.; Goodrich, S.; Higgins, L.C., eds. 2016. A Utah Flora. Fifth Edition, revised. Provo, UT: Brigham Young University. 990 p.
Wood, D.M.; del Moral, R. 1987. Mechanisms of early primary succession in subalpine habitats on Mount St. Helens. Ecology. 68(4): 780-790.
Young, S.A.; Schrumpf, B.; Amberson, E. 2003. The Association of Official Seed Certifying Agencies (AOSCA) native plant connection. Moline, IL: AOSCA. 9 p.
Zika, P.F.; Alverson, E.R.; Wagner, W.H.; Wagner, F.S. 2002. Botrychium hesperium in the Wallowa Mountains of Oregon. American Fern Journal. 92(3): 239-240.
How to Cite
Gucker, Corey L.; Shaw Nancy L. 2021. Annual agoseris (Agoseris aurantiaca [Hook.] Greene). In: Gucker, C.L.; Shaw, N.L., eds. Western forbs: Biology, ecology, and use in restoration. Reno, NV: Great Basin Fire Science Exchange. Online: http://westernforbs.org/species/orange-agoseris-agoseris-aurantiaca/