Authorship

Gucker, Corey; Shaw, Nancy

Publication Date

October 2020

Table of Contents (collapsible)

    Nomenclature

    Nineleaf biscuitroot (Lomatium triternatum [Pursh] J.M. Coult. & Rose) belongs to the Apiaceae family, Apioideae subfamily, and Peucedaneae tribe (Harborne and Williams 1972; Albach et al. 2001).

    Family

    Apiaceae – Carrot family

    Genus

    Lomatium

    Species

    triternatum

    NRCS Plant Code

    LOTR2 (USDA NRCS 2019).

    Subtaxa

    As of 2020, the following nineleaf biscuitroot varieties were recognized: Lomatium triternatum var. anomalumL. t. var. brevifoliumL. t. var. macrocarpumL. t. var. triternatum (ITIS 2020; USDA NRCS 2019).

    Synonyms

    L. t. subsp. platycarpumL. t. var. alatum (ITIS 2020).

    Common Names

    Nineleaf biscuitroot, broadnineleaf biscuitroot, narrowleaf biscuitroot, narrowleaf desert-parsley, nineleaf lomatium, and ternate lomatium.

    Chromosome Number

    2n = 22 (Munz and Keck 1973; Constance et al. 1976; Hickman 1993; Welsh et al. 2016). 2n = 22 was reported for var. anomalum (Constance et al. 1976) and brevifolium (Rice et al. 2015). 2n = 22, 44 were reported for varieties macrocarpum and triternatum (Munz and Keck 1973; Hickman 1993).

    Hybridization

    No hybridization has been documented for the Lomatium genus (Hitchcock and Cronquist 2018).

    Distribution

    Nineleaf biscuitroot is most common in the northwestern US states of Washington, Oregon, and Idaho (USDA FS 1937; Soltis et al. 1997) and also occurs in the southern portions of British Columbia and Alberta, Canada. The species range covers both sides of the Cascade Range and Sierra Nevada Mountains, all but the northernmost and easternmost parts of Montana, all but the easternmost part of Wyoming, and northern California, northern Nevada, eastern Utah, western Colorado, and northwestern New Mexico (BONAP 2017).

    Habitat And Plant Associations

    Nineleaf biscuitroot is adapted to intense sun exposure (Fig. 1). Plants have leaves with very narrow blades that resist water loss with excessive radiation. It is common in open terrain, rocky sites, meadows, and mountain sites with high sun exposure (Daubenmire 1975a). Nineleaf biscuitroot occurs in dry to moist plains, foothills, lower and mid-montane locations, and is occasional at high-elevation sites averaging 90 or more frost-free days (USDA FS 1937; Lambert 2005a; Tilley et al. 2010; Pavek et al. 2012). It occurs at sites receiving 8 to 20 in (203–508 mm) of annual precipitation that are typically moist in spring but dry by early summer (Taylor 1992; Skinner 2007).

    Several nineleaf biscuitroot plants in flowering growing with sagebrush and grasses.

    Figure 1. Nineleaf biscuitroot growing in a dry sagebrush community in Oregon. Photo: Bureau of Land Management (BLM) OR014, Seeds of Success (SOS).

    Nineleaf biscuitroot commonly occurs in grassland, sagebrush (Artemisia spp.) steppe, mountain shrub, pinyon-juniper (PinusJuniperus spp.), ponderosa pine (P. ponderosa), and alpine communities (USDA FS 1937; Munz and Keck 1973; Stevens and Monsen 2004; Tilley et al. 2010). It is especially common in the antelope bitterbrush-bluebunch wheatgrass (Purshia tridentataPseudoroegneria spicata) rangeland cover type in the Pacific Northwest and in the basin big sagebrush (Artemisia tridentata subsp. tridentata), mountain big sagebrush (A. t. subsp. vaseyana), threetip sagebrush (A. tripartita), and low sagebrush (A. arbuscula) rangeland cover types in the Great Basin (Shiflet 1994).

    Forests and woodlands. Nineleaf biscuitroot is an indicator of dry forest and woodland communities in the West (Fig. 2). In Oregon and California, it occurred in western juniper (J. occidentalis) woodlands where soil moisture availability was limited by layers of hard clay and duripan (Driscoll 1964; Miller et al. 2000). On pumice soils in central Oregon, nineleaf biscuitroot was most common in the driest, lowest elevation ponderosa pine/antelope bitterbrush communities (Dyrness and Youngberg 1966). In western Montana, nineleaf biscuitroot was an indicator of xeric subalpine larch (Larix lyallii) stands based on the species composition of 127 stands found near timberline at cold, rocky sites (Arno and Habeck 1972).

    A dense stand of nineleaf biscuitroot growing near a stream and a conifer forest.

    Figure 2. Nineleaf biscuitroot growing in a montane forest opening in Oregon. Photo: BLM OR014, SOS.

    Shrublands. Sagebrush shrublands are common habitat for nineleaf biscuitroot (Fig. 3). In Oregon and Idaho, nineleaf biscuitroot occurred in low sagebrush, mountain big sagebrush, threetip sagebrush, and antelope bitterbrush shrublands (Tisdale et al. 1965; Dealy and Geist 1978; Doescher et al. 1986; Huntly et al. 2011). On Silver Lake winter range in south-central Oregon, nineleaf biscuitroot was a prominent forb in low sagebrush at 5,200 ft (1,585 m) where annual precipitation averaged 15 in (380 mm) (Dealy and Geist 1978). Nineleaf biscuitroot was a principal forb in threetip sagebrush/Idaho fescue (Festuca idahoensis) vegetation on north and east slopes and in threetip sagebrush/Idaho fescue/bluebunch wheatgrass vegetation on south and west slopes at Craters of the Moon National Monument, Idaho. The threetip sagebrush community occupied moister, cooler sites with relatively deep, fertile soils (Tisdale et al. 1965).

    Grasslands. Although nineleaf biscuitroot is common in bluebunch wheatgrass grasslands (Fig. 4), but is more abundant in Idaho fescue grasslands (Tisdale 1979). In western Montana, nineleaf biscuitroot averaged 1.8% cover in Idaho fescue and 0.4% cover in bluebunch wheatgrass grasslands (Mitchell 1957). In Power County, Idaho, nineleaf biscuitroot frequency and production was lower in grasslands on north- (0.9 lbs/ac [1 kg/ha]; 1% freq) than south-facing slopes (7 lbs/ac [8 kg/ha]; 32% freq) with extremely stony, silt loam soils (Shiflet 1975). Nineleaf biscuitroot was also associated with rough fescue (F. campestris)-bluebunch wheatgrass and rough fescue-Idaho fescue communities in western Montana (Mueggler and Stewart 1980). In the Bitterroot Mountains, nineleaf biscuitroot was common in high-elevation grassland balds (>6000 ft [1,830 m]). The balds are considered climax vegetation and occur on south-facing slopes within the spruce-fir (PiceaAbies spp.) zone where soils are coarsely fractured and excessive summer moisture deficiencies restrict tree establishment (Root and Habeck 1972).

    Dense patches of nineleaf biscuitroot growing between sagebrush shrubs in a dry conifer forest opening.

    Figure 3. Nineleaf biscuitroot growing in an upper elevation sagebrush community abutting a montane forest in Oregon. Photo: BLM OR014, SOS.

    A handful of nineleaf biscuitroot plants growing in a mix of grasses with scattered sagebrush.

    Figure 4. Nineleaf biscuitroot growing with bluebunch wheatgrass near Boise, ID. Photo: US Forest Service, Rocky Mountain Research Station (USFS RMRS).

    Elevation

    Nineleaf biscuitroot is most common at low and mid-elevations but also occurs at elevations up to 10,000 ft (3,000 m) (USDA FS 1937). In California, nineleaf biscuitroot often occurs at elevations below 5,000 ft (1,500 m) (Munz and Keck 1973), but ranges from 660 to 6,600 ft (200–2,000 m) (Hickman 1993). In Utah, the elevation range is 4,300 to 9,500 ft (1,300–2,900 m) (Tilley et al. 2010). High elevation occurrences of 9,500 ft (2,900 m) were reported from Utah and 10,000 ft (3,000 m) from Wyoming (USDA FS 1937).

    Soils

    Nineleaf biscuitroot commonly grows in relatively neutral (6.5–8.8 pH), well-drained or dry, rocky soils (USDA FS 1937; Stevens and Monsen 2004; Tilley et al. 2010). Vegetation and habitat studies report nineleaf biscuitroot in sandy to clay loam soils 8 to 39 in (20–100 cm) deep (Driscoll 1964; Dealy and Geist 1978; Doescher et al. 1986; Bryant et al. 2013). Although rare in clay-rich soils (Darrach et al. 2010), clay or hardpan layers beneath coarser soil layers are common (Driscoll 1964; Miller et al. 2000).

    Several studies provide additional information about the soil characteristics in nineleaf biscuitroot habitats. In an evaluation of vegetation along a moisture gradient in the Siskiyou Mountains of Oregon and California, nineleaf biscuitroot abundance was greater at xeric, low-elevation sites with serpentine soils (Whittaker 1960). Near Craters of the Moon National Monument, Idaho, nineleaf biscuitroot was a principal forb in deep fertile soils dominated by threetip sagebrush/Idaho fescue vegetation (Tisdale et al. 1965). In western Montana, nineleaf biscuitroot abundance was significantly greater (P < 0.05) in low nitrogen than high nitrogen plots when control, thinned, burned, and thinned and burned ponderosa pine stands were compared (Gundale et al. 2006).

    In Oregon and northeastern California, nineleaf biscuitroot was found in sandy loams, loams, and clayey soils. In the High Desert and Klamath Ecological Provinces of southeastern Oregon and northeastern California, nineleaf biscuitroot occurred in low sagebrush/Sandberg bluegrass (Poa secunda) communities on clayey soils 8 to 24 in (20–60 cm) deep underlain by fractured basalt. It was also common in mountain big sagebrush/Thurber’s needlegrass (Achnatherum thurberianum) vegetation on loam soils 14 to 26 in (36–67 cm) deep underlain by hardpan (Miller et al. 2000). On a semi-isolated plateau between the Crooked and Deschutes river canyons in central Oregon, nineleaf biscuitroot occurred in western juniper woodlands in non-rocky, loess, and very fine sand soils with a dense clay or stony clay loam layer at 8 to 15 in (20–38 cm) deep. Soils averaged 1.5% organic matter, 0.08% total nitrogen, and pH 6.5 to 8 (Driscoll 1964). In Silver Lake winter range, nineleaf biscuitroot grew with low sagebrush and antelope bitterbrush in soils with fines, gravels, and rocks up to 8 in (20 cm) in size. Soils included basalt, andesite, pumice, ash, and obsidian (Dealy and Geist 1978).

    Resource acquisition. In field experiments, nineleaf biscuitroot nitrogen uptake was monitored throughout the spring by nitrogen type in a sagebrush community in eastern Oregon. Nitrogen (NH4+, NO3) was injected into soil around nineleaf biscuitroot at various depths in the spring (25 April and 23 May at 0.8–2.8 in [2–7 cm] and 6.7–8.7 in [17–22 cm] deep). In April, nineleaf biscuitroot captured about 20% of NH4+ and 50% of NO3 provided and less than 20% of both nitrogen types in May. Plants captured about the same amount of each type of nitrogen type (a little less than 20%) at 0.8–2.8 in (2–7 cm) deep, but much more NO3 (50%) than NH4+ (10%) at 6.7 to 8.7 in (17–22 cm) deep (James 2009).

    Description

    Nineleaf biscuitroot is a long-lived perennial forb (Tilley et al. 2010). Plants are 8 to 32 in (20–80 cm) tall with slender, hollow, unbranched stems and leaves that are primarily or entirely basal (Munz and Keck 1973; Patterson et al. 1985; LBJWC 2020). Plants grow from a long, thick, somewhat woody taproot ranging from slender to massive and reaching several feet deep (USDA FS 1937; Hickman 1993). Five-year-old plants growing in seed production plots had 11 in (28 cm) long taproots weighing 0.28 lb (125 g) (Tilley et al. 2010).

    Stems and leaves are green to blue-green, nearly hairless or with fine, soft, appressed hairs, and smell like celery (Fig. 5; Welsh et al. 2016; Luna et al. 2018). Leaves are ternately to ternate-pinnately compound with 9 to 21 narrowly linear, well-separated leaflets or segments (USDA FS 1937; Tilley et al. 2010; Welsh et al. 2016; Luna et al. 2018). Leaf petioles are up to 8 in (20 cm) long with dilated bases forming a sheath around the stem. Leaflets range from 0.4 to 5 in (1–13 cm) long and 0.4 to 0.6 in (1–1.5 cm) wide (Tilley et al. 2010; Welsh et al. 2016).

    Inflorescences are compound umbels (Fig. 5; Pavek et al. 2012). The umbels have 4 to 20 unequal rays ranging up to 3.5 in (9 cm) long, which make the umbel irregular or lop-sided (USDA FS 1937; Luna et al. 2018). Rays are topped with umbellets, which contain numerous tiny yellow flowers (Fig. 6; Tilley et al. 2010). Umbellets on the longer rays support most of the bisexual flowers with some male flowers, while those on shorter rays support mainly male flowers (Welsh et al. 2016; Hitchcock and Cronquist 2018). Individual flowers have five bright yellow stamens and five yellow petals, which fade to white with age (Welsh et al. 2016). Flowers produce schizocarps, which are fruits with two seeds (mericarps). Schizocarps are dry, oblong to elliptical, and 0.2 to 0.6 in (6–15 mm) long by 0.2 to 0.4 in wide (4–11 mm) wide (Patterson et al. 1985). Mericarps separate along their midlines when mature. Mericarps are strongly flattened, unnotched at the base and apex with five equal ribs on the dorsal side and thin wings along the sides (USDA FS 1937; Luna et al. 2018).

    Single nineleaf biscuitroot plant with fringy leaves and terminal flowers on the tallest stems.

    Figure 5. Nineleaf biscuitroot plant with highly dissected leaves and umbellets with small yellow flowers in compound umbels. Photo: BLM OR014 SOS.

    Nineleaf biscuitroot compound umbel with >6 rays of unequal lengths.

    Figure 6. Nineleaf biscuitroot’s compound umbel with the typical unequal ray lengths and umbellets in full bloom. Photo: BLM ID931 SOS.

    Varieties. Nineleaf biscuitroot varieties in some locations are poorly defined (Hickman 1993) and in other locations are in taxonomic flux (Lesica and Kittelson 2013; Sosa et al. 2015). In California, variety macrocarpum is a stout plant that produces 0.3 to 0.8-in (8-20 mm) long fruits and occurs at 660 to 5,000 ft (200–1,500 m) elevation (Munz and Keck 1973; Hickman 1993).

    Researchers found no intermediate forms when studying varieties anomalum and triternatum that co-occurred near Hot Springs, Montana, which suggested no interbreeding and that the varieties should be recognized as two species (Lesica and Kittelson 2013). The varieties grew within 16 to 98 ft (5–30 m) of each other. Variety anomalum leaflets (6–11 mm) were wider than those of variety triternatum (2–4 mm) with no overlap. Mericarps of variety anomalum were larger (8.6 mm long, 4.8 mm wide) than those of variety triternatum (7 mm long, 4.2 mm wide) with little overlap. Specific leaf area of variety anomalum was about half that of variety triternatum. Variety anomalum occurred in swales dominated by tall, leafy forbs, while variety triternatum was most common on more xeric ridgetops and slopes dominated by sparse bluebunch wheatgrass tussocks (Lesica and Kittelson 2013).

    Reproduction

    Nineleaf biscuitroot reproduces from seed (Fig. 7; USDA FS 1937). Plants produce male and bisexual flowers (Schlessman 1982). Bisexual flowers are commonly found on the longer outer rays of the umbels and male flowers on the shorter inner umbel rays (Welsh et al. 2016; Hitchcock and Cronquist 2018). The proportion of male flowers increases in later flowering umbels (Skinner 2007).

    Fruit set for biscuitroot species depends on pollination. Fernleaf biscuitroot (L. dissectum) fruit set was 60 times greater for flowers freely visited by pollinators than those protected from pollinators (Cane et al. 2020). Production of flowers fluctuates greatly from year to year within and between populations (Gilmartin 1982). Bisexual flowers produce the schizocarps that later split into two flat mericarps. A single umbel can produce up to several hundred mericarps (Thompson 1985).

    Phenology

    Nineleaf biscuitroot begins growing in early spring (Tilley et al. 2010). Plants generally flower in April and May, and seeds are typically mature in June and July (Stevens and Monsen 2004; Lesica and Kittelson 2010; BLM SOS 2017). Nineleaf biscuitroot phenology was monitored for 2 years in big sagebrush/bluebunch wheatgrass in southern British Columbia on steep slopes at 1,800 to 2,460 ft (550–750 m) where annual precipitation averaged 7 in (188 mm). Plant growth began in March, flowering in April, and plants were in full flower in May. Seed set and shatter occurred in May and June, and plants were cured in July (Pitt and Wikeem 1990).

    Pollination

    Native ground nesting bees are important nineleaf biscuitroot pollinators, especially mining bees (Andrena or Micandrena spp.) (Cane and Love 2016). In a survey of bees and their floral associations at nine sites in Idaho, Oregon, and Wyoming, an average of 10 bees/100 plants were collected from late April to June sweeps of 1,309 nineleaf biscuitroot plants. Bees represented eight genera, predominantly pollen specialists: 51% mini-mining bees (Micrandrena spp., an Andrena subgenus), 32% nonsocial, ground-nesting, mining bees, 0.7% small carpenter bees (Ceratina spp.), 4% sweat bees (Halictus spp.), 8% sweat bees (Lasioglossum spp.), 2% cuckoo bees (Nomada spp.), 2% confluent miner bees (Panurginus spp.), and 0.7% mason bees (Osmia spp.) (Cane and Love 2016). In another survey, 149 bees representing 20 species were collected from nineleaf biscuitroot flowers at 12 western US sites. Mining bees were most common (A. microchloraA. chlorogasterA. gordoni primarily) (Cane et al. 2020).

    Nineleaf biscuitroot provides habitat for mining bee species in fire-prone areas (Cane et al. 2020). Fires typically occur when nineleaf biscuitroot is dormant and likely to survive burning. Mining bees nest underground for one season and are active in the spring but deep underground (7.5 in [19 cm]) by summer. Seasonality of nineleaf biscuitroot and its mining bee pollinators should, therefore, survive wildfires relatively unscathed and be ready to reproduce the following spring. Consequently, when present before a fire, this pollinator relationship should recover quickly. However, without good knowledge of pre-fire communities or fire conditions, restoration is sometimes initiated even though natural regeneration may occur. If restoration includes creation of deep furrows, the apical taproot meristems of nineleaf biscuitroot can be damaged, setting back natural or assisted regeneration (Cane et al. 2020).

    Ecology

    Nineleaf biscuitroot is a long-lived (20+ yrs), mid- to late-seral perennial (Walker and Shaw 2005; Tilley et al. 2010). It produces a well-developed taproot, and its above-ground growth structure and phenology make it drought, grazing, and fire tolerant (Daubenmire 1975a; Stevens and Monsen 2004; Skinner et al. 2005).

    Seed And Seedling Ecology

     Seed predation is common for nineleaf biscuitroot (Thompson 1985). In a grassland in eastern Washington, seeds were taken by small mammals and beetles. When 54 mixed piles of fernleaf biscuitroot and nineleaf biscuitroot (5 of each) were monitored, more fernleaf biscuitroot than nineleaf biscuitroot seeds were taken, and fernleaf biscuitroot seeds were taken significantly faster than nineleaf biscuitroot seeds (< 0.0001). Preference may have been influenced by nineleaf biscuitroot seed size which was 10 to 20% larger than nineleaf biscuitroot (Thompson 1985).

    Above-ground seedling growth is slow, and plants produce only a few leaves in their first year because energy is devoted to taproot development (Ogle et al. 2012).

    Nineleaf biscuitroot compound umbel with schizocarps that are oval shaped, with pale yellow wings, and pink ribs.

    Figure 7. A single nineleaf biscuitroot compound umbel with mature fruits (schizocarps). Photo: BLM ID931, SOS.

    Disturbance Ecology

    Nineleaf biscuitroot is somewhat disturbance tolerant and generally more common in mid- to late-seral communities that are in good ecological condition with a diversity of other forb species. In canyon grasslands south of Lewiston, Idaho, nineleaf biscuitroot cover increased with increasing seral stage. Cover averaged 0.2% in annual-dominated, early seral communities, 0.4% in mid-seral communities, and 0.5% in perennial-dominated, late-seral communities. Nineleaf biscuitroot was negatively associated (P < 0.05) with yellow starthistle (Centaurea solstitialis), the dominant species in early seral communities (Robins 2001). In Clarkston, Washington, nineleaf biscuitroot was absent from agricultural fields abandoned for 1 to 12 years, rare in older fields abandoned for 39 to 52 years but had a frequency of up to 62% in never cultivated steppe vegetation (Daubenmire 1975b). In Benton County, Washington, nineleaf biscuitroot occurred in a 52-year-old field where sampling was concentrated at field edges closest to native, undisturbed vegetation (Simmons and Rickard 2002).

    Nineleaf biscuitroot populations produced seed in the ash fallout from the eruption of Mt. St. Helens in Washington (May 1980). These plants showed no difference from those evaluated the year before the eruption in number of leaves produced, sex ratio, or proportion of flowering individuals (Gilmartin 1982).

    Fire. Nineleaf biscuitroot typically survives fire and its abundance is often greater on burned than unburned sites (Daubenmire 1975b; Busse et al. 2000; Riegel et al. 2006). Cover of nineleaf biscuitroot was not significantly different on unburned, low-severity, or high-severity burned plots in the first 3 years following fire in ponderosa pine forests on the Coeur d’Alene Indian Reservation (Armour et al. 1984). Following an August fire in a sagebrush-grassland near Pocatello, Idaho, nineleaf biscuitroot cover was significantly greater on burned plots that were not seeded with a mixture of nonnative grasses and forbs. In the first postfire growing season (May), nineleaf biscuitroot cover was 1.0% in seeded and 1.8% in unseeded plots (Ratzlaff and Anderson 1995).

    Following summer fires in Idaho and Washington, nineleaf biscuitroot abundance was greater on burned than unburned plots and increased with time since fire (Daubenmire 1975b; Gucker 2004). Frequency of nineleaf biscuitroot was greater on burned than unburned bluebunch wheatgrass-Sandberg bluegrass communities near Clarkston, Washington. Frequency was about six times greater on burned than unburned plots in the second postfire year, about four times in the fourth postfire year, and about equal by the eighth postfire year (Daubenmire 1975b). Following a low-severity, summer fire in northern Idaho, nineleaf biscuitroot cover increases occurred on both burned and unburned plots but changes from pre-fire levels were greater on burned plots (Table1; Gucker 2004).

    Table 1. Nineleaf biscuitroot cover (%) after a low-severity fire in bluebunch wheatgrass-Sandberg bluegrass-arrowleaf balsamroot vegetation south of Lewiston, Idaho (Gucker 2004).

    Post fire year

    2 yrs before fire

    1 yr before fire

    2 yrs post-fire

    3 yrs post-fire

    ————-% cover————-

    Unburned

    0.16

    0.42

    1.58

    1.04

    Burned

    0.40

    0.39

    2.20

    4.02

    Although nineleaf biscuitroot abundance was often greater on burned than unburned plots, that was not the case following an intense August fire in Gambel oak (Quercus gambelii) and big sagebrush/bluebunch wheatgrass vegetation near Midway, Utah (Table 2). In the first postfire year, nineleaf biscuitroot cover and frequency were much lower on burned than unburned plots in both vegetation types (Table 2; Poreda and Wullstein 1994).

    Table 2. Frequency and cover (%) of nineleaf biscuitroot in the first postfire year following a severe summer fire near Midway, Utah (Poreda and Wullstein 1994).

    Vegetation type

    Unburned

    Burned

    Frequency

    Cover

    Frequency

    Cover

    ———————%———————

    Big sagebrush/bluebunch wheatgrass

    27.3

    0.91

    4.9

    0.02

    Gambel oak

    35.3

    0.77

    1.8

    0.01

    High levels of seedling establishment were reported for one combination of seeding rate and heat produced in a field experiment at Morgan Prairie, in Thurston County, Washington. Plots (10 ft² [1 m²]) were burned at low (547 °F [286 °C]), moderate (648 °F [342 °C]), and high surface temperatures (1,047 °F [564 °C]) and then seeded with a mix of 10 native species at rates of 0, 50, 100, and 200 seeds/m² (5, 9, and 19/ft²). Nineleaf biscuitroot recruitment was nearly 5 seedlings/ft² (50/m²) at moderately burned plots seeded at 200 seeds/m² (19/ft²), which was much greater than any other treatment combination. Recruitment of nineleaf biscuitroot was almost zero on unburned plots and less than 0.5 seedlings/ft² (5/m²) on low or high temperature burns (Reagan 2014).

     

    Wildlife And Livestock Use

    Nineleaf biscuitroot provides important spring forage to native ungulates, domestic grazers, bears, small mammals, greater sage-grouse, and insects (Gullion 1960; Mueggler and Stewart 1980; Knight and Blanchard 1983; Cox 1989; Pyle 1993; Landis and Savoie 2018). It is considered valuable forage because of its large stature and high-production levels (Tilley et al. 2010). Elk (Cervis canadensis), deer (Odocoileus spp.), pronghorn (Antilocapra americana), and bighorn sheep (Ovis canadensis) feed on nineleaf biscuitroot throughout its range (Gullion 1960). Forage is considered good for elk and deer, fair to good for sheep, and poor to fair for cattle (Mueggler and Stewart 1980).

    Mammals. Amounts and seasons of use of nineleaf biscuitroot were reported for several mammals in the West. In south-central Oregon, mule deer (Odocoileus hemionus) fed on flower heads and flower stalks in meadows in a lodgepole pine forest (Stuth and Winward 1977). Mountain goats (Oreamnos americanus) fed on nineleaf biscuitroot in June in western Montana’s Swan Mountains (Chadwick 1974). On the Desert Game Range, in Clark County, Nevada, bighorn sheep fed on nineleaf biscuitroot in July (Hall cited in Gullion 1960). Domestic sheep readily consume its leaves, flowers, and green seeds (USDA FS 1937). On elk winter range in Cow Creek, Montana, nineleaf biscuitroot represented a trace of cattle diets in late July and 3% of diets in August (Gordon 1968). In Yellowstone National Park, nineleaf biscuitroot was 1% of grizzly bear (Ursus arctos) diets by volume from scat analysis (Knight and Blanchard 1983). Northern pocket gophers (Thomomys talpoides) feed on nineleaf biscuitroot and it is a highly preferred food source. At Lawrence Memorial Grassland Preserve, north-central Oregon, all 9 nineleaf biscuitroot roots in monitored tunnels were taken. At this site, many plants were rooted within large grass clumps, which may offer protection from predation (Cox 1989).

    Birds. Nineleaf biscuitroot foliage and insect visitors are important to greater sage-grouse (Centrocercus urophasianus). Leaves are eaten in early spring, and the flowers support invertebrates, which are a critical food source for greater sage-grouse chicks (Rhodes et al. 2010; Tilley et al. 2010; Pennington et al. 2016; Luna et al. 2018). At the Hart Mountain National Antelope Refuge in Lake County, Oregon, nineleaf biscuitroot was recovered from the crops of greater sage-grouse chicks collected from brood-rearing habitat (Pyle 1993).

    Insects. Early spring green-up and flowering make nineleaf biscuitroot important for spring pollinators, especially pollen-collecting mining bees (Tilley et al. 2010). Nineleaf biscuitroot is a host species for swallowtail butterflies (Papilio zelicaon and P. indra) (James and Nunnallee 2011; Landis and Savoie 2018) and a nectar source for Puget blue (Icaricia icarioides blackmorei) and other butterflies (LaBar and Schultz 2012).

    Many mining bee species have been collected from nineleaf biscuitroot including Andrena angustitarsataA. auricomaA. forbesiiA. fulvinigraA. hippotesA. merriamiA. oligolegeA. pallidifoveaA. saliciflorisA. scurraA. striatifronsA. quintiliformis, and A. wheeleri (La Berge 1973; Ribble 1974; Laberge and Ribble 1975; LaBerge 1989).

    Ethnobotany

    Indigenous people living in the western US and Canada used nineleaf biscuitroot for food, medicine, ceremonies, and luck. Roots, leaves, and flowers were eaten but roots were the staple (Johnston 1970; Reveal et al. 1999). Roots, leaves, flowers, and fruits were used medicinally.

    Nineleaf biscuitroot roots were eaten raw, roasted, or dried for later use by many Indigenous groups (Johnston 1970; Marshall 1977; Turner et al. 1980; Reveal et al. 1999; Moerman 2003). The Atsugewi of northern California, cooked the roots in earth ovens (Moerman 2003). The Okanagan-Colville, Blackfeet, and other tribes in Montana baked biscuits and cakes from flour made from the roots (Patterson et al. 1985; Moerman 2003). The Blackfeet used nineleaf biscuitroot flowers to make pemmican (Moerman 2003). The Okanagan-Colville used dried flowers and upper leaves to add a celery-like flavor to meats, stews, and salads. Sanpoil and Nespelem tribes ate roots raw or pit-cooked and laid leaves and roots on top of cooking baskets to add flavor (Turner et al. 1980).

    Medicinally, nineleaf biscuitroot was used by Indigenous people to treat respiratory illnesses. The Blackfeet considered nineleaf biscuitroot to be a cure-all (Moerman 2003). Chewed roots were made into a spray and believed to penetrate and heal the body at the point of pain. An infusion of roots was used to treat chest troubles, and fruits were chewed by long-distance runners to avoid side aches (Moerman 2003). The Okanagan-Colville used an infusion of flowers and upper leaves for colds and sore throats (Turner et al. 1980).

    There were several other miscellaneous uses for nineleaf biscuitroot. It was carried by Indian men as a love charm (Van Allen Murphey 1990). Seeds were stuffed in a porcupine foot that was tied in the hair of young Blackfeet girls for good luck (Moerman 2003). Seeds were also used during the tanning of animal pelts to control the odor. The Paiute believed they could summon cold winds by breaking nineleaf biscuitroot stems or roots (Moerman 2003).

    Horticulture

    Nineleaf biscuitroot is an attractive, long-lived, drought-tolerant plant that would be useful in low-maintenance landscapes such as campgrounds, roadsides, or rest areas where native and adapted. Bright yellow flowers and interesting brown striped seeds grow on upright plants with unusual leaves. Its use has been encouraged for native landscaping in the Palouse region of Idaho and Washington where it thrives with little added water or fertilizer (Skinner et al. 2005).

    Revegetation Use

    Nineleaf biscuitroot is recommended for pollinator and wildlife habitat conservation plantings and provides fair soil stabilization where native and adapted (Shaw and Monsen 1983; Tilley et al. 2013; Dumroese et al. 2016). It grows well in mountain big sagebrush, pinyon-juniper, aspen (Populus tremuloides), and subalpine zones receiving annual precipitation of 12 to 20 in (305–508 mm) (Tilley et al. 2013; Ogle et al. 2019). Once established it is competitive with weeds and tolerant of drought, grazing, and fire (Daubenmire 1975a; Stevens and Monsen 2004; Skinner et al. 2005; Tilley et al. 2010).

    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 partners to first select appropriate species and seed sources and then properly collect, grow, certify, clean, store, and distribute seed for restoration (PCA 2015).

    Developing a seed supply begins with seed collection from native stands. Collection sites are determined by current or projected revegetation requirements and goals. 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).

    Seed Sourcing

    Because empirical seed zones are not currently available for nineleaf biscuitroot, 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 8, Omernik Level III Ecoregions (Omernik 1987) overlay the provisional seed 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.

    Occurrence Map

    Figure 8. Distribution of nineleaf biscuitroot (black circles) based on geo-referenced herbarium specimens and observational data from 1881-2016 (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

    Releases

    As of 2020, there were no nineleaf biscuitroot germplasm releases.

    Wildland Seed Collection

    Nineleaf biscuitroot seeds are typically ready for harvest when the inflorescence is dry and fruits are tan, which is generally in late June or July (Skinner 2007). Because many lomatium species have similar looking seeds, it is best to collect from plants that still have leaves to aid identification (Camp and Sanderson 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 (Young et al. 2020; UCIA 2015). 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 Field 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.

    Collection Timing

    At maturity nineleaf biscuitroot seeds are dry, papery, tan to brown, and easily dislodged from the inflorescence (Fig. 9). This typically occurs 5 to 8 weeks after flowering in late June or July. (Parkinson and DeBolt 2005; Camp and Sanderson 2007; Skinner 2007; BLM SOS 2017). Seed ripens uniformly within individual umbels but ripening can vary among umbels on the same plant or on different plants (Skinner 2007).

    The Bureau of Land Management’s Seeds of Success crews made 48 collections of nineleaf biscuitroot over 9 years between 2000 to 2016 (USDI BLM SOS 2017). Harvests were made in Washington, Oregon, California, Nevada, Utah, Idaho, and Wyoming. Sixty seven percent of the collections were made in July, and 25% were made in June. The earliest collection was made on May 22, 2013, from 3,340 ft (1,020 m) in elevation in Malheur County, Oregon. The latest harvest was made on September 2, 2013, from 5,700 ft (1,740 m) in elevation in Lassen County, California. In the single year with the greatest number of collections (19 harvests in 2010), the earliest was June 21 from 2,968 ft (905 m) in elevation in Ada County, Idaho, and the latest was July 27 at 6,570 ft (2,000 m) in elevation in Uinta County, Wyoming (USDI BLM SOS 2017).

    Nineleaf biscuitroot plant with dry staw colored stems and schizocarps and mericarps. Some mericarps dispersing.

    Figure 9. Nineleaf biscuitroot with mature seed dispersing. Photo: BLM CA350 SOS.

    Collection Methods

    Wildland seed is easily hand collected by stripping or shaking umbels over a container or by clipping the inflorescences (Fig. 10; Jorgensen and Stevens 2004; Parkinson and DeBolt 2005; Skinner 2007). Seed detaches readily from the umbellets. Collections made by shaking ripe inflorescences over a container are very clean (Ogle et al. 2012).

    Woman collecting nineleaf biscuitroot seed on a slope with grasses and sagebrush.

    Figure 10. Collecting wildland nineleaf biscuitroot seed in a sagebrush community. Photo: USFS RMRS.

    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 nineleaf biscuitroot.

    Collection Rates

    Hand-stripping is easy but can be moderately time consuming, depending on stand conditions (variability in seed ripening, plant distribution) if a large quantity of seed is required (Fig. 11; Camp and Sanderson 2007).

    Post-Collection Management

    Seed should be kept in a dry, shaded place until it can be moved to a controlled short-term storage environment. Short-term storage should be in a dry, cool location inaccessible to rodents or other seed predators. If insects are suspected in any collection, seed should be frozen for 48 hours or treated with an appropriate insecticide. The more plant material in the harvest, the more ventilation and drying a seed lot may need (Gold n.d.; Parkinson and DeBolt 2005; Hay and Probert 2011). Camp and Sanderson (2007) recommend drying nineleaf biscuitroot at room temperature for 2 to 4 weeks before cleaning.

    Seed Cleaning

    Seed can be cleaned to high purity with minimal air screening (Ogle et al. 2012). Seeds must be handled carefully as they are fragile and brittle. Small collections can be carefully hand-rubbed to free seeds from the inflorescence then cleaned using an air column separator. Large collections require cleaning with air screen equipment (Skinner 2007).

    An umbellet with 4 schizocarps, 3 detached schizocarps, and a single mericarp. The schizocarps and mericarps are straw colored, have narrow papery wings, and raised ribs. Mericarp has a midline where was attached to another mericarp.

    Figure 11. Nineleaf biscuitroot fruits (winged schizocarps with ribs) and a mericarp (center) with the commissure visible. Photo: USFS RMRS.

    A small seed lot collected from the Columbia Basin was cleaned by hand rubbing inflorescences, then using an air column separator, and finally shaking seed through a standard soil testing sieve (3.5 meshes/in). Some chaff remained following this process (Camp and Sanderson 2007).

    A larger, hand-collected seed lot (2.5 lbs [1.1 kg]) was cleaned at the USFS Bend Seed Extractory. Seed was processed using a Westrup Model LA-H laboratory brush machine (Corvallis, OR), with a #20 mantel and medium speed. Seed was then air-screened using an office Clipper (Bluffton, IN) with a 1/12 round top and 7 ¾ cross slot bottom screen. Seed was rescreened using a 1/12 top round screen, 18 bottom round screen, medium speed, and medium air (Barner 2007).

    Seed Storage

    Nineleaf biscuitroot seed retains its viability after 3 to 4 years of storage (Stevens and Monsen 2004). Recommended storage conditions range from 33 to 40 °F (0.6–4 °C) and 35 to 44% relative humidity (Barner 2007; Camp and Sanderson 2007; Skinner 2007). Nineleaf biscuitroot seed viability was 92% after 18 months of storage at room temperature in a glass container (Parkinson and DeBolt 2005). Viability of multiple collections ranged from 44 to 95% after dry seed was stored 5 months at 68 °F (20 °C) at the Royal Botanical Gardens Kew. Pre-storage viability was not reported (RGB Kew 2017).

    Seed Testing

    Seed fill can be evaluated be viewing mericarps over a light table (Parkinson and DeBolt 2005). There is no Association of Official Seed Analysts (AOSA) rule for testing germination of nineleaf biscuitroot. Vankus (2006) recommends testing germination at temperatures of 59 to 77 °F (15–25 °C). First seed count is made after 7 days and the final count at 28 days. Fresh seed requires 28 to 63 days of cold stratification at 37 to 41 °F (3–5 °C). Older seed may germinate during stratification (Vankus 2006).

    Viability Testing

    The following tetrazolium chloride (TZ) testing procedure can be used to evaluate viability of nineleaf biscuitroot seed. Barely cover seed with water and soak it overnight at room temperature, then cut the seed longitudinally to remove about one quarter of the mericarp. Discard the quarter sections and soak the rest of the mericarp for 6 hours or overnight in a 1% TZ solution. Staining is evaluated by making a longitudinal cut at the center of the seed through the embryo. Normal live embryos stain pink or reddish. Dead embryos are unstained, partially stained, or very darkly stained (Hurd n.d.).

    Germination Biology

    Nineleaf biscuitroot produces seed with immature embryos that require a long period (9+ weeks) of cold stratification to develop and germinate (Shaw and DeBolt 2005; Skinner 2007; RBG Kew 2017). Germination occurs at low temperatures, light is not required, and gibberellic acid fails as a cold (< 50 °F [10 °C]) stratification substitute (Shaw and DeBolt 2005). In trials conducted at the NRCS Pullman Plant Materials Center, nineleaf biscuitroot failed to germinate after 30 days of cold stratification, but seed sown outdoors in November emerged in late March (Skinner 2007).

    Several studies report on the necessity of cold stratification for germination. Jorgensen and Wilson (2004) reported 50% germination after 100 days in the dark at 34 to 38 °F (1–3 °C) and a little less than 70% after 250 days (Jorgensen and Wilson 2004). Parkinson and DeBolt (2005) achieved 93% germination when imbibed seeds were chilled at 39 °F (4 °C) in the dark for 60 days. Germination began after 36 days of stratification, and most seeds germinated within 49 days. Seeds were soaked for 24 hours before the cold stratification treatment (Parkinson and DeBolt 2005). The Royal Botanical Gardens Kew obtained 100% germination for 19 seeds placed on 1% agar, treated to 8 hours of light and 16 hours of dark at 41 °F (5 °C) for 258 days (RBG Kew 2017).

    Drake et al. (1998) and Drake and Ewing (n.d.) reported no germination (0%) with 6 weeks stratification and 6.7% germination following 12 weeks of cold stratification at 36 to 43 °F (2–6 °C). Germination results were the same for seed germinated outdoors (48–65 °F [9–18 °C]) or in a greenhouse (65–85 °F [18–29 °C]) following the stratification treatments. Seed was 63% viable based on TZ testing.

    Wildland Seed Yield And Quality

    Post-cleaning seed yield and quality of nineleaf biscuitroot seed lots collected in the Intermountain region are provided in Table 3 (USFS BSE 2017). Results indicate that fresh nineleaf biscuitroot seed purity, fill, and viability can be highly variable. Most other estimates of seeds/lb reported elsewhere in the literature fall within those reported in Table 3 (37,130–63,812 seeds/lb [81,856–140,679 seeds/kg) (Lambert 2005a; Skinner 2007; USFS GBNPP 2014; RBG Kew 2017), but some reports exceeded those values (90,500 seeds/lb [199,515 seeds/kg]) (Wiese et al. 2012) and 113,500 seeds/lb [250,220 seeds/kg]) (Camp and Sanderson 2007).

    Table 3. Seed yield and quality of nineleaf biscuitroot 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

    Mean

    Range

    Samples (no.)

    Bulk weight (lbs)

    1.5

    0.11–5.6

    26

    Clean weight (lbs)

    0.7

    0.05–3.0

    26

    Clean-out ratio

    0.5

    0.10–0.8

    26

    Purity (%)

    92

    66–99

    26

    Fill (%)¹

    87

    50–98

    26

    Viability (%)²

    88

    59–98

    25

    Seeds/lb

    50,464

    32,000–87,000

    26

    Pure live seeds/lb

    38,740

    14,807–73,602

    25

    ¹100 seed X-ray test
    ²Tetrazolium chloride test

    Marketing Standards

    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. The following marketing standards were suggested for nineleaf biscuitroot seed: acceptable purity 75%, viability 70%, and germination 70% (Walker and Shaw 2005).

    Agricultural Seed Production

    Trials for growing nineleaf biscuitroot for seed production were conducted at Oregon State University’s Malheur Experimental Station (OSU MES) (Shock et al. 2016) and the NRCS Aberdeen Plant Materials Center (Ogle et al. 2012). At OSU MES, production plots were planted with seed collected near Lakeview, Oregon. Plants began flowering and produced some seed in their second year, but seed yields were low until year 3 (Shock et al. 2018). Seed was harvested for 11 years at OSU MES, where they found that seed production increased with irrigation in most years (see Irrigation section) (Shock et al. 2016, 2018). Seed production plots at the Aberdeen PMC did not flower or produce seed until year 4 (Ogle et al. 2012).

    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 (Young et al. 2020; UCIA 2015).

    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.

    Site Preparation

    Seeding is best in firm, weed-free seedbeds. Seed production plots that were used to test preemergence herbicides at OSU MES were seeded at a rate of 30 pure live seed (PLS)/ft of row (98 PLS/m) on December 1, 2009. Seed was placed on the soil surface then covered with a thin layer of sawdust (0.18 oz/ft [0.6 g/m] of row) and row cover until April 7, 2010 (Shock et al. 2011).

    Weed Management

    Weed control is necessary for good seed yields (Tilley et al. 2010). Weed control can be achieved using weed barrier fabric, cultivation, and hand roguing (Ogle et al. 2012; Shock et al. 2016). Because nineleaf biscuitroot goes dormant early in the growing season, plots can be treated later in the growing season without compromising seed production (St. John and Ogle 2011).

    Trials at OSU MES evaluated emergence of nineleaf biscuitroot following preemergent herbicide treatments (Shock et al. 2011). Most herbicide treatments had minimal effect on emergence, which was only about 25% regardless of treatment (Table 4). Preemergent herbicides were applied in late November using a CO2 sprayer at 30 PSI (Shock et al. 2011).

    Table 4. Emergence of nineleaf biscuitroot in spring 2010 following preemergence herbicide treatments in fall 2009 in experimental plots at Oregon State University’s Malheur Experiment Station, Ontario, OR (Shock et al. 2011).

    Herbicide applied Nov 30, 2009

    Application rate
    (lb ai/ac)

    Emergence (%)

    Untreated

    NA

    26.0

    Bensulide

    5.0

    26.5

    Pronamide

    1.0

    17.2

    Trifluralin

    0.375

    24.2

    Pendimethalin

    0.75

    24.3

    Benefin

    1.2

    26.3

    Dimethenamid

    0.656

    21.2

    Linuron

    0.5

    24.0

    *Providing charcoal protection to the seed with treatments of pendimethalin had a significantly detrimental effect on nineleaf biscuitroot emergence.

    Yields of nineleaf biscuitroot seed were not significantly different with most annual post-emergent herbicide treatments (Table 5), except the first year’s treatment of bromoxynil, which significantly reduced seed yields compared to untreated plants. Herbicides were applied in the spring for 4 consecutive years. Seed production was very low in 2009, the reason for this was not reported. Seed yields represent small, one-row plots without borders and may not be representative (Shock et al. 2011).

    Table 5. Seed yield of nineleaf biscuitroot following the second, third, and fourth years of post-emergent herbicide treatments in experimental plots seeded in November 2005 at Oregon State University’s Malheur Experiment Station, Ontario, OR (Shock et al. 2011).

    Herbicide

    Application rate
    (lb ia/ac)

    Year 2

    Year 3

    Year 4

    3/12/2008

    3/20/2009

    4/7/2010

    Seed yields (lbs/ac)

    Control

    NA

    981.5a

    32.6a

    475.3a

    Bromoxynil

    0.125

    187.9b

    20.8a

    462.8a

    Oxyfluorfen

    0.125

    820.8a

    28.2a

    421.6a

    Clethodim

    0.094

    1,062.1a

    38.2a

    449.1a

    Pendimethalin

    1.0

    922.7a

    28.6a

    488.7a

    Prometryn

    0.8

    1,069.8a

    27.8a

    449.1a

    Dimethenamid

    0.656

    987.0a

    26.6a

    413.4a

    Linuron

    0.5

    888.1a

    31.2a

    533.3a

    At the Aberdeen PMC, nineleaf biscuitroot was direct seeded in November 2005 in weed barrier fabric. Weeds were controlled with wick applications of glyphosate and hand weeding while nineleaf biscuitroot was establishing and actively growing. In June 2007, survival was 71%, but no plants flowered. By early July, plants were dormant, and plots were sprayed with glyphosate. In 2008 and 2009, plants produced seed in July. Seed production was nearly twice as much in 2009 as in 2008. Plots were glyphosate treated in November in 2008 and 2009. Better seed production in 2009 than in 2008 was due to plant age and not related to weed treatments (St. John and Ogle 2011).

    Seeding

    Nineleaf biscuitroot was seeded in late October 2005 at rates of 20 to 30 PLS/linear ft (66–98 PLS/m) in seed production plots at OSU MES. It was seeded in silt loams using a custom, small-plot, grain drill with disk openers. Seed was planted 0.5 in (1.3 cm) deep (Shock et al. 2016, 2018). Seeds were placed on the soil surface, then buried with a thin layer of sawdust. A long period of cold stratification is needed for seed to germinate, and in spring 2006, stands were considered excellent (Shock et al. 2016, 2018). Seedlings failed to emerge when seed was cold stratified for 1 month prior to early spring seeding on March 3 (Shock et al. 2018).

    At the Aberdeen PMC, nineleaf biscuitroot was fall seeded in rows at a rate of 25 PLS/ft (82 PLS/m) of row or into weed barrier fabric at 18 to 24-in (40–60 cm) spacings (Ogle et al. 2012). In the establishment of a common garden near Boise State University, nineleaf biscuitroot was seeded 0.4 to 0.8 in (1–2 cm) deep in holes burned into weed barrier cloth in fall 2004. Emergence began in late February 2005 when a high percentage of holes had seedlings (Shaw and DeBolt 2005).

    Establishment And Growth

    In its first year of growth, nineleaf biscuitroot produces few leaves and devotes energy to taproot development (Ogle et al. 2012). In the early seedling stage, nineleaf biscuitroot can be mistaken for a grass due to its linear cotyledons (Fig. 12; Pavek et al. 2012). Plants are considered established after a year and flower after 2 to 4 years of growth (Ogle et al. 2012; Shock et al. 2016). Once established, nineleaf biscuitroot is a strong perennial when protected from rodents (Shock et al. 2018). Plants green up, flower, and set seed early in the growing season (Table 6).

    Many seedlings emerging from soil, some with mericarp still attached at bright green cotelydon tips. Seedlings red at the base.

    Figure 12. Nineleaf biscuitroot seedlings emerging from a southern Idaho planting. Photo: USFS RMRS.

    Table 6.Timing of flowering and seed harvest dates for nineleaf biscuitroot seed production fields over a period of 9 years at Oregon State University’s Malheur Experiment Station, Ontario, OR (Shock et al. 2016). Seed yields are provided in Table 7 (Shock et al. 2016, 2018).

    Year

    Flower start

    Flower peak

    Flower end

    Harvest

    ———————– Date ———————–

    2007

    25 Apr

    1 Jun

    29 Jun, 16 Jul

    2008

    25 Apr

    5 Jun

    3 Jul

    2009

    10 Apr

    7 May

    1 Jun

    26 Jun

    2010

    25 Apr

    15 Jun

    22 Jul

    2011

    30 Apr

    23 May

    15 Jun

    26 Jul

    2012

    12 Apr

    17 May

    6 Jun

    21 Jun

    2013

    18 Apr

    10 May

    4 Jun

    2014

    7 Apr

    29 Apr

    2 May

    4 Jun

    2015

    10 Apr

    28 Apr

    20 May

    7 or 15 Jun

    2016

    11 Apr

    28 Apr

    20 May

    15 Jun

    2017

    24 Apr

    15 May

    30 May

    27 Jun

    Irrigation

    Trials conducted at OSU MES (Fig. 13) found that plant size and seed yield of nineleaf biscuitroot were significantly greater with irrigation (Table 7) (Shock et al. 2018). Irrigation trials delivered 0, 4, or 8 in (101 or 203 mm) of water in March 2006 to plots that were seeded in late October 2005. Irrigation treatments were delivered four times at about 2-week intervals beginning at the time of bud formation and flowering. Irrigation was delivered through drip tape buried 12 in (30 cm) deep, so as not to affect flowering and to limit germination of weed seed (Shock et al. 2016). In most years, nineleaf biscuitroot seed yields increased significantly with irrigation (P < 0.05). Seed yield was maximized by 4 in (101 mm) or more of water in cool, wet years and by 8 in (203 mm) of water in warm, dry years (Shock et al. 2016). Seed yields exhibited a quadratic response to irrigation from 2008 to 2012 and a linear response to irrigation in 2007, 2013, 2014, and 2015 (Shock et al. 2016, 2018). Accumulated growing degree hours were higher than average in 2006, 2007, and 2013 to 2016. High accumulated growing degree hours in 2015 were probably the reason for early harvest dates (Shock et al. 2018).

    Pollinator Management

    Nineleaf biscuitroot flowers attract and are primarily pollinated by mining bees that nest underground and live for 1 year. Seed production plots may attract mining bees from nearby populations of spring-blooming floral hosts (Salicaceae, Rosaceae, Brassicaceae). When grown near towns or apiaries, nineleaf biscuitroot flowers are visited by honeybees suggesting field pollination could be improved by providing hives (Cane et al. 2020).

    Table 7. Nineleaf biscuitroot seed yields (lbs/ac) with and without irrigation for 11 years of varied precipitation and growing conditions at Oregon State University’s Malheur Experiment Station, Ontario, OR (Shock et al. 2016, 2018).

    Year

    Precipitation levels

    Growing degree hrs (50-86 °F) +/- the 25-yr mean

    Added irrigation

    0 in

    4 in

    8 in

    Yield (lbs/ac)

    2007

    Below normal

    +66

    2.3a

    17.5ab

    26.7b

    2008

    Below normal

    +199

    195a

    1,061b

    1,387b

    2009

    Above normal

    -120

    182a

    780b

    676b

    2010

    Above normal

    0

    1,637a

    2,830b

    3,195c

    2011

    Above normal

    -236

    1,983a

    2,625b

    2,028a

    2012

    Normal

    -351

    239a

    603b

    733b

    2013

    Below normal

    +21

    154a

    734b

    1,051b

    2014

    Below normal

    +112

    241a

    897b

    1,497c

    2015

    Normal

    +403

    403a

    441a

    955b

    2016

    Normal

    +251

    395a

    476ab

    639b

    2017

    Above normal

    -11

    933a

    949a

    1,266b

    11-yr ave

    9.8 in (fall + winter + spring)

    1,207 hrs

    579a

    1,038b

    1,211c

    Pest Management

    Insects and mammals can affect nineleaf biscuitroot growth and reproduction (see Seed and Seedling Ecology and Wildlife and Livestock Use sections). Nineleaf biscuitroot is also host to various fungi: Asperisporium peucedaniCercosporella herpotrichoidesFusicladium peucedaniLeptosphaeria conoideaNodulosphaeria modestaPhoma lomatiae, and Puccinia jonesii (Farr and Rossman 2017). Effects or impacts of fungi on nineleaf biscuitroot growth or survival were not reported.

    Seed Harvesting

    At OSU MES, nineleaf biscuitroot was harvested by hand and cleaned using an Office Clipper (Bluffton, IN) (Shock et al. 2015). At the Aberdeen PMC harvested nineleaf biscuitroot seed was harvested using vacuum type or flail vac harvesters. These harvests were easily air screened to nearly 100% purity (Ogle et al. 2012). Nineleaf biscuitroot seed harvested using the Jet Harvester developed by the Aberdeen PMC was much more efficient than hand harvesting, and the seed required little post-harvest cleaning (Tilley and Bair 2011).

    Large nineleaf biscuitroot plants with multiple umbels growing in rows.

    Figure 13. Nineleaf biscuitroot irrigation trial at OSU MES. Photo: USFS RMRS.

    Seed Yields And Stand Life

    Nineleaf biscuitroot was harvested for 11 years at OSU MES (Shock et al. 2018). Averaged over the years, seed yield was maximized at 1,213 lbs/ac (1,360 kg/ha) with 12.4 in of water (irrigation + spring precipitation) (Shock et al. 2018).

    Regression analyses estimated that seed yields were greatest with 4 to 8 in (102–210 mm) of irrigation, depending on the year (Table 8; Shock et al. 2016). Seed yields showed a quadratic response to irrigation plus spring precipitation from 2008 through 2013. In 2007, and 2014 to 2016, seed yield showed a positive linear response to irrigation plus spring precipitation. In 2017, seed yields were not responsive to irrigation, likely because of heavy winter and spring precipitation (Shock et al. 2018).

    Table 8. Regression analysis for fernleaf biscuitroot seed yield (y, lbs/ac) (in response to irrigation plus spring precipitation (x, mm/season) using equation y=a + b∙x + c∙x²) in 2007-2017 and the 11-year average. Study conducted at Oregon State University’s Malheur Experiment Station in Ontario, OR. Only those years where irrigation resulted in significantly greater seed yields are shown below (Shock et al. 2018).

    Year

    Max yield (lb/ac)

    Water applied plus spring ppt for max yield (in/season)

    Spring ppt (in)

    2007

    28

    9.9

    1.9

    2008

    1,390

    9.8

    1.4

    2009

    824

    9.5

    4.1

    2010

    3,196

    12.1

    4.3

    2011

    2,623

    8.8

    4.8

    2012

    734

    10.8

    2.6

    2013

    1,115

    11.6

    0.9

    2014

    1,509

    9.7

    1.7

    2015

    875

    11.2

    3.2

    2016

    624

    10.2

    2.2

    Ave

    1,213

    12.4

    2.9

    *2017 seed yield was not responsive to irrigation, probably because of heavy winter and spring precipitation (Table 7).

    Nursery Practice

    Nineleaf biscuitroot plants were grown successfully outdoors and in greenhouses (Parkinson and DeBolt 2005; Camp and Sanderson 2007), but plants started outdoors that were moved to a greenhouse died at the Aberdeen PMC. Causes of mortality were thought to be due to temperature changes or root pathogens (Skinner 2007).

    Parkinson and DeBolt (2005) planted germinating seed 0.4 in (1 cm) deep in Styrofoam cone-tainers filled with equal parts peat and vermiculite. Cone-tainers were kept at a constant 81 °F (27 °C) in the greenhouse and watered when saturation levels fell below 80%. Small amounts of seedling fertilizer were added periodically. True leaves appeared after 14 days. Seedling survival was 53% after 77 days in the greenhouse (Parkinson and DeBolt 2005).

    Camp and Sanderson (2007) planted nineleaf biscuitroot in a soil mix of 50% peat, 35% perlite, and 15% sand in early November. Seeds were lightly covered with sand and kept outdoors. Germination was good and began in late February. Seedlings developed 3 to 4 leaves in the spring. Planting beds were top dressed with NPK 14-14-14 fertilizer. Good stands re-emerged the following spring and about 5% of the plants bloomed. Outdoor beds were not watered or fertilized in the second year (Camp and Sanderson 2007).

    Nineleaf biscuitroot plants were grown successfully outdoors at the Pullman PMC. Seed was collected from the Paradise Creek Drainage near Pullman, Washington. Seed was sown in Ray Leach Super cell cone-tainers (10 cu in) filled with Sunshine #4 soil mix. Seeds were covered lightly with soil and a thin layer of coarse grit to prevent seeds from floating when watered. Cone-tainers were watered deeply and placed outside. Additional watering occurred only during dry spells. Seedling emergence began in March with warm daytime temperatures and occurred over a period of 3 to 4 weeks. Seedlings were watered as needed and fertilized once a week with a complete, water-soluble fertilizer. Seedlings were moved to a lath house in early June, where they were only watered to prevent complete drying. Plants overwintered in a lath house, where they were protected from rodents and covered with mulch or foam sheets in extremely cold temperatures. A few plants flowered in the year following outplanting but most required 2 years in the field to produce flowers and seed (Skinner 2007).

    Wildland Seeding And Planting

    Nineleaf biscuitroot is recommended for restoration or revegetation in sagebrush, pinyon-juniper, and aspen zones where precipitation averages 8 to 25 in (203–635 mm)/year (Shaw and Monsen 1983; Walker and Shaw 2005; Ogle et al. 2011). Nineleaf biscuitroot provides good pollinator habitat and fair soil stabilization (Shaw and Monsen 1983; Ogle et al. 2011; Eldredge et al. 2013). Although nineleaf biscuitroot has a slow initial growth rate, it is competitive with weeds once established and persists as a mid- to late-seral species (Walker and Shaw 2005; Tilley et al. 2010).

    Fall seeding is necessary for seed stratification and germination. Seed can be broadcast or drill seeded 0.1 to 0.2 in (0.3–0.6 cm) deep as a mix or separately (Shaw and Monsen 1983; Stevens and Monsen 2004; Tilley et al. 2010). The pure seeding rate is 20 to 24 PLS lbs/ac (22–27 kg/ha) (Ogle et al. 2011). In wildland restoration efforts, nineleaf biscuitroot was seeded in mixes at a rate of 0.2 to 1.5 lbs/ac (0.2–1.7 kg/ha) (Cosgriff et al. 2004; Lambert 2005b; Applestein et al. 2018). Newly seeded sites should be protected from grazing for at least 2 years (Stevens and Monsen 2004).

    Transplants and root material were established successfully in field evaluations at the Aberdeen PMC. Transplants grown for a full season and overwintered before planting were very hardy and established at almost 100% when weeds were controlled (Tilley et al. 2010). At Canaday Farm in Spring Creek, Nevada, root pieces 2 to 8 in (5–20 cm) long had good emergence when field planted in the spring of 2010. By April 13, 2011, multiple leaflets and flowering stems had emerged from most roots. The orientation and size of the roots in the planting hole did not impact emergence. Plants generally produced aboveground growth proportional to the size of the transplanted rootstock. Height of the plants, which included flowering stems, was typically less than 8 in (20 cm). Survival through the first spring was good and some plants produced limited amounts of seed (Jensen and Stettler 2012).

    Acknowledgements

    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. We give great thanks to the chapter reviewers: Alexis Gibson, Program Manager for the Society for Ecological Restoration and Matt Lavin, Professor of Plant Sciences and Plant Pathology at Montana State University.

    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.

    Literature Cited

    Albach, D.C.; Soltis, P.S.; Soltis, D.E.; Olmstead, R.G. 2001. Phylogenetic analysis of Asterids based on sequences of four genes. Annals of the Missouri Botanical Garden. 88(2): 163-212.

    Applestein, C.; Germino, M.J.; Pilliod, D.S.; Fisk, M.R.; Arkle, R.S. 2018. Appropriate sample sizes for monitoring burned pastures in sagebrush steppe: How many plots are enough, and can one size fit all? Rangeland Ecology and Management. 71(6): 721-726.

    Armour, C.D.; Bunting, S.C.; Neuenschwander, L.F. 1984. Fire intensity effects on the understory in ponderosa pine forests. Journal of Range Management. 37(1): 44-49.

    Arno, S.F.; Habeck, J.R. 1972. Ecology of alpine larch (Larix lyallii Parl.) in the Pacific Northwest. Ecological Monographs. 42(4): 417-450.

    Barner, J. 2007. Propagation protocol for production of propagules (seeds) Lomatium triternatum (Pursh.) Coult. & Rose seeds. Native Plant Network. U.S. Department of Agriculture, Forest Service, National Center for Reforestation, Nurseries, and Genetic Resources. http://npn.rngr.net/propagation/protocols [Accessed 2020 July 1].

    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.

    Biota of North America Program [BONAP]. 2017. http://www.bonap.org/.

    Bower, A.D.; St. Clair, J.B.; Erickson, V. 2014. Generalized provisional seed zones for native plants. Ecological Applications. 24(5): 913-919.

    Bryant, J.A.M.; Reynecky, B.K.; Brown, R.L. 2013. Geology and topography effects on exotic plant distribution in a semiarid Mima mound prairie in eastern Washington. Northwest Science. 87(1): 12-23.

    Busse, M.D.; Simon, S.A.; Riegel, G.M. 2000. Tree-growth and understory responses to low-severity prescribed burning in thinned Pinus ponderosa forests of central Oregon. Forest Science. 46(2): 258-268.

    Camp, P.; Sanderson, J. 2007. Seed collection, propagation and reintroduction of native wildflowers in the Columbia Basin. Wenatchee, WA: U.S. Department of the Interior, Bureau of Land Management, Wenatchee Field Office. 32 p.

    Cane, J.H.; Love, B. 2016. Floral guilds of bees in sagebrush steppe: Comparing bee usage of wildflowers available for postfire restoration. Natural Areas Journal. 36(4): 377-391.

    Cane, J.H.; Weber, M.; Love, B.C. 2020. Self-compatibility in Lomatium dissectum (Apiaceae) and the diverse Andrena bees that dominate regional Lomatium pollinator faunas. Western North American Naturalist. 80(1): 1-10.

    Chadwick, D.H. 1974. Mountain goat ecology: Logging relationships in the Bunker Creek drainage of western Montana. Missoula, MT: University of Montana. Thesis. 262 p.

    Consortium of Pacific Northwest Herbaria [CPNWH]. 2017. Seattle, WA: University of Washington Herbarium, Burke Museum of Natural History and Culture. http://www.pnwherbaria.org/index.php

    Constance, L.; Chuang, T.I.; Bell, C.R. 1976. Chromosome numbers in Umbelliferae. V. American Journal of Botany. 63(5): 608-625.

    Cox, G.W. 1989. Early summer diet and food preferences of northern pocket gophers in north central Oregon. Northwest Science. 63(3): 77-82.

    Darrach, M.; Thie, K.K.; Wilson, B.L.; Brainerd, R.E.; Otting, N. 2010. Lomatium tamanitchii (Apiaceae) a new species from Oregon and Washington state, USA. Madrono. 57(3): 203-208.

    Daubenmire, R. 1975a. Floristic plant geography of eastern Washington and northern Idaho. Journal of Biogeography. 2(1): 1-18.

    Daubenmire, R. 1975b. Plant succession on abandoned fields, and fire influences, in a steppe area in southeastern Washington. Northwest Science. 49(1): 36-48.

    Dealy, J.E.; Geist, J.M. 1978. Conflicting vegetational indicators on some central Oregon scablands. Journal of Range Management. 31(1): 56-59.

    Doescher, P.S.; Miller, R.F.; Swanson, S.R.; Winward, A.H. 1986. Identification of the Artemisia tridentata ssp. wyomingensis/Festuca idahoensis habitat type in eastern Oregon. Northwest Science. 60(1): 55-60.

    Drake, D.; Ewing, K. nd. Germination requirements of 32 native Washington prairie species. Seattle, WA: The Nature Conservancy: 181-190.

    Drake, D.; Ewing, K.; Dunn, P. 1998. Techniques to promote germination of seed from Puget Sound prairies. Restoration and Management Notes. 16(1): 33-40.

    Driscoll, R.S. 1964. A relict area in the central Oregon Juniper zone. Ecology. 45(2): 345-353.

    Dumroese, R.K.; Luna, T.; Pinto, J.R.; Landis, T.D. 2016. Forbs: Foundation for restoration of monarch butterflies, other pollinators, and greater sage-grouse in the western United States. Native Plants Journal. 36(4): 499-511.

    Dyrness, C.T.; Youngberg, C.T. 1966. Soil-vegetation relationships within the ponderosa pine type in the central Oregon pumice region. Ecology. 47(1): 122-138.

    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/.

    Gilmartin, A.J. 1982. Effects on Lomatium triternatum of the 1980 ash fallout from Mt. St. Helens. Madrono. 29(4): 270-271.

    Gold, K. n.d. Post-harvest handling of seed collections. Tech. Information Sheet 04. UK: Royal Botanic Gardens Kew and Millennium Seed Bank Partnership. 4 p.

    Gordon, F.A. 1968. Range relationships of elk and cattle on elk winter range, Cow Creek, Montana. Bozeman, MT: Montana State University. Thesis. 40 p.

    Gucker, C.L. 2004. Canyon grassland vegetation changes following the Maloney Creek wildfire. Moscow, ID: University of Idaho. Thesis. 29 p.

    Gullion, G.W. 1960. The ecology of Gambel’s quail in Nevada and the arid southwest. Ecology. 41(3): 518-536.

    Gundale, M.J.; Metlen, K.L.; Fiedler, C.E.; DeLuca, T.H. 2006. Nitrogen spatial heterogeneity influences diversity following restoration in a ponderosa pine forest, Montana. Ecological Applications. 16(2): 479-489.

    Harborne, J.B.; Williams, C.A. 1972. Flavonoid patterns in the fruits of the umbelliferae. Phytochemistry. 11(5): 1741-1750.

    Hay F.R.; Probert, R.J. 2011. Chapter 20: Collecting and handling seeds in the field. In: Guarino, L.; Ramanatha, V.; Goldberg, E. Collecting plant genetic diversity: Technical Guidelines-2011 update. Rome, Italy: Bioversity International. 33 p.

    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. Second Ed. Giblin, D.E.; Legler, B.S.; Zika, P.F.; Olmstead, R.G., eds. Seattle, WA: University of Washington Press. 882 p.

    Huntly, N.; Bangert, R.; Hanser, S.E. 2011. Native and exotic plants of fragments of sagebrush steppe produced by geomorphic processes versus land use. Plant Ecology. 212(9): 1549-1561.

    Hurd, E.G. n.d. Tetrazolium test for selected species. Unpublished report on file with the USDA Forest Service, Rocky Mountain Research Station, Boise, ID. 30 p.

    ITIS Database. 2020. Integrated Taxonomic Information System. Available: http://www.itis.gov/index.html

    James, D.G.; Nunnallee, D. 2011. Life histories of Cascadia butterflies. Corvallis, OR: Oregon State University Press. 447 p.

    James, J. 2009. Linking species and functional group diversity to resource capture and invasion resistance in Great Basin plant communities. In: Shaw, N.; Pellant, M., eds. Great Basin Native Plant Selection and Increase Project: 2008 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 106-112.

    Jensen, S.; Stettler, J. 2012. Applying provisional seed zones to Great Basin forb production and cultural practice notes. In: Shaw, N.; Pellant, M., eds. Great Basin Native Plant Project: 2011 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 70-84.

    Johnston, A. 1970. Blackfoot Indian utilization of the flora of the northwestern Great Plains. Economic Botany. 24(3): 301-324.

    Jorgensen, K.R.; Stevens, R. 2004. Seed collection, cleaning, and storage. In: Monsen, S.B.; Stevens, R.; Shaw, N.L., comps. Restoring western ranges and wildlands, vol. 3. Gen. Tech. Rep. RMRS-GTR-136-vol-3. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 699-716.

    Jorgensen, K.R.; Stevens, R. 2004b. Seed germination. In: Monsen, S.B.; Stevens, R.; Shaw, N.L., comps. Restoring western ranges and wildlands, vol. 3. Gen. Tech. Rep. RMRS-GTR-136-vol-3. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 723-732.

    Knight, R.R.; Blanchard, B.M. 1983. Yellowstone grizzly bear investigations: Annual report of the Interagency Study Team, 1982. Washington, DC: U.S. Department of the Interior. National Park Service. 45 p.

    LaBar, C.C.; Schultz, C.B. 2012. Investigating the role of herbicides in controlling invasive grasses in prairie habitats: Effects on non-target butterflies. Natural Areas Journal. 32(2): 177-189.

    LaBerge, W.E. 1973. A revision of the bees of the genus Andrena of the Western Hemisphere. Part VI. Subgenus Trachandrena. Transactions of the American Entomological Society. 99(3): 235-371.

    LaBerge, W.E. 1989. A revision of the bees of the genus Andrena of the Western Hemisphere. Part XIII. Subgenera Simandrena and Taeniandrena. Transactions of the American Entomological Society. 115(1): 1-56.

    Laberge, W.E.; Ribble, D.W. 1975. A revision of the bees of the genus Andrena of the Western Hemisphere. Part VII. Subgenus Euandrena. Transactions of the American Entomological Society. 101(3): 371-446.

    Lady Bird Johnson Wildflower Center [LBJWC]. 2020. Lomatium triternatum (Pursh) J.M. Coult. & Rose. Native Plant Database. Austin, TX: Lady Bird Johnson Wildflower Center. https://www.wildflower.org/plants-main [Accessed 2020 July 20].

    Lambert, S. 2005. Guidebook to the seeds of native and non-native grasses, forbs and shrubs of the Great Basin. Boise, ID: U.S. Department of the Interior, Bureau of Land Management, Idaho State Office. 136 p.

    Lambert, S.M. 2005b. Seeding considerations in restoring big sagebrush habitat. In: Shaw, N.L.; Pellant, M.; Monsen, S.B., eds. Sage-grouse habitat restoration symposium proceedings; 2001 June 4-7; Boise, ID. Proc. RMRS-P-38. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 75-80.

    Landis, T.D.; Savoie, S. 2018. Using native plants to create pollinator habitat in southwest Oregon: Lessons learned. Native Plants Journal. 19(1): 27-39.

    Lesica, P.; Kittelson, P.M. 2010. Precipitation and temperature are associated with advanced flowering phenology in a semi-arid grassland. Journal of Arid Environments. 74(9): 1013-1017.

    Lesica, P.; Kittelson, P.M. 2013. Morphological and ecological segregation of two sympatric Lomatium triternatum (Apiaceae) varieties in Montana. Madroño. 60(3): 211-216.

    Luna, T.; Mousseaux, M.R.; Dumroese, R.K. 2018. Common native forbs of the northern Great Basin important for greater sage-grouse. Gen. Tech. Rep. RMRS-GTR-387. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station; Portland, OR: U.S. Department of the Interior, Bureau of Land Management, Oregon-Washington Region. 76 p.

    Marshall, A.G. 1977. Nez Perce social groups: An ecological interpretation. Pullman, WA: Washington State University. Thesis. 194 p.

    Miller, R.F.; Svejcar, T.J.; Rose, J.A. 2000. Impacts of western juniper on plant community composition and structure. Journal of Range Management. 53(6): 574-585.

    Mitchell, W.W. 1957. An ecological study of the grasslands in the region of Missoula, Montana. Missoula, MT: University of Montana. Thesis. 111 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

    Mueggler, W.F.; Stewart, W.L. 1980. Grassland and shrubland habitat types of western Montana. Gen. Tech. Rep. INT-66. Ogden, UT: U.S. Department of Agriculture, Forest Service, Intermountain Forest and Range Experiment Station. 154 p.

    Munz, P.A.; Keck, D.D. 1973. A California flora and supplement. Berkeley, CA: University of California Press. 1905 p.

    Ogle, D.; Pavek, P.; Fleenor, R.; Stannard, M.; Dring, T.; Cane, J.; Fullen, K.; St. John, L.; Tilley, D. 2011. Plants for pollinators in the Inland Northwest. Tech. Note 2B. Boise, ID: U.S. Department of Agriculture, Natural Resources Conservation Service. 65 p.

    Ogle, D.; Shaw, N.; Cane, J.; St. John, L.; Tilley, D. 2012. Propagation protocol for production of propagules (seeds) Lomatium triternatum (Pursh.) Coult. & Rose seeds. Native Plant Network. U.S. Department of Agriculture, Forest Service, National Center for Reforestation, Nurseries, and Genetic Resources. http://npn.rngr.net/propagation/protocols [Accessed 2020 July 1].

    Ogle, D.; Tilley, D.; Cane, J.; St. John, L.; Fullen, K.; Stannard, M.; Pavek, P. 2019. Plants for pollinators in the Intermountain West. Plant Materials Tech. Note 2A. Boise, ID: U.S. Department of Agriculture, Natural Resources Conservation Service. 40 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.

    Parkinson, H.; DeBolt, A. 2005. Propagation protocol for production of container (plugs) Lomatium triternatum (Pursh.) Coult. & Rose seeds. Native Plant Network. U.S. Department of Agriculture, Forest Service, National Center for Reforestation, Nurseries, and Genetic Resources. http://npn.rngr.net/propagation/protocols [Accessed 2020 July 1].

    Patterson, P.A.; Neiman, K.E.; Tonn, J. 1985. Field guide to forest plants of northern Idaho. Gen. Tech. Rep. RMRS-GTR-118. Ogden, UT: U.S. Department of Agriculture, Forest Service. 246 p.

    Pavek, P.; Erhardt, B.; Heekin, T.; Old, R. 2012. Forb seedling identification guide for the Inland Northwest: Native, introduced, invasive, and noxious species. Pullman, WA: U.S. Department of Agriculture, Natural Resources Conservation Service, Pullman Plant Materials Center. 144 p.

    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.

    Pitt, M.D.; Wikeem, B.M. 1990. Phenological patterns and adaptations in an Artemisia/Agropyron plant community. Journal of Range Management. 43(4): 350-358.

    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.

    Poreda, S.F.; Wullstein, L.H. 1994. Vegetation recovery following fire in an oakbrush vegetation mosaic. Great Basin Naturalist. 54(4): 380-383.

    Pyle, W.H. 1992. Response of brood-rearing habitat of sage grouse to prescribed burning in Oregon. Corvallis, OR: Oregon State University. Thesis. 47 p.

    Ratzlaff, T.D.; Anderson, J.E. 1995. Vegetal recovery following wildfire in seeded and unseeded sagebrush steppe. Journal of Range Management. 48(5): 386-391.

    Reagan, K. 2014. Restoration of native plant communities: An examination of seed limitation and microsite limitation in the Garry oak ecosystem. Seattle, WA: University of Washington. Dissertation. 82 p.

    Reveal J.L., Moulton G.E., Schuyler A.E. 1999. The Lewis and Clark collections of vascular plants: names, types, and comments. Proceedings of the Academy of Natural Sciences of Philadelphia. 149: 1-64.

    Rhodes, E.C.; Bates, J.D.; Sharp, R.N.; Davies, K.W. 2010. Fire effects on cover and dietary resources of sage-grouse habitat. The Journal of Wildlife Management. 74(4): 755-764.

    Ribble, D.W. 1974. A revision of the bees of the genus Andrena of the Western Hemisphere subgenus Scaphandrena. American Entomological Society. 100(2): 101-189.

    Rice, A.; Glick, L.; Abadi, S.; Einhorn, M.; Kopelman, N.M.; Salman-Minkov, A.; Mayzel, J.; Chay, O.; Mayrose, I. 2015. The Chromosome Counts Database (CCDB) – A community resource of plant chromosome numbers. New Phytologist. 206(1): 19-26.

    Richardson, B.; Kilkenny, F.; St. Clair, B.; Stevenson-Molnar, N. 2020. Climate Smart Restoration Tool. https://climaterestorationtool.org/csrt/

    Riegel, G.M.; Miller, R.F.; Skinner, C.N.; Smith, S.E. 2006. Northeastern Plateaus bioregion. In: Sugihara, N.G.; van Wagtendonk, J.W.; Shaffer, K.E.; Fites-Kaufman, J.; Thode, A., eds. Fire in California’s ecosystems. Berkeley, CA: University of California Press: 225-263.

    Robins, S.S. 2001. Effects of perennial plant competition on the invasibility of canyon grassland communities by Centaurea solstitialis. Moscow, ID: University of Idaho. Thesis. 48 p.

    Root, R.A.; Habeck, J.R. 1972. A study of high elevational grassland communities in western Montana. The American Midland Naturalist. 87(1): 109-121.

    Royal Botanic Gardens Kew [RBG Kew]. 2017. Seed Information Database (SID). Version 7.1. http://data.kew.org/sid/

    Schlessman, M.A. 1982. Expression of andromonoecy and pollination of tuberous Lomatiums (Umbelliferae). Systematic Botany. 7(2): 134-149.

    SEINet – Regional Networks of North American Herbaria Steering Committee [SEINet]. 2017. SEINet Regional Networks of North American Herbaria. https://symbiota.org/seinet/

    Shaw, N.; DeBolt, A. 2005. Development of germination protocols, seed weight, purity, and seed conditioning/cleaning protocols for Great Basin grasses and forbs. In: Shaw, N.L.; Pellant, M., eds. Great Basin Native Plant Selection and Increase Project: 2004 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 9-13.

    Shaw, N.; Monsen, S.B. 1983. Nonleguminous forbs for rangeland sites. In: Monsen, S.B; Shaw, N. Managing Intermountain rangelands-Improvement of range and wildlife habitats; 1981 September 15-17; Twin Falls, ID; 1982 June 22-24; Elko, NV. Gen. Tech. Rep. INT-157. Ogden, UT: U.S. Department of Agriculture, Forest Service, Intermountain Forest and Range Experiment Station: 123-131.

    Shiflet, T.N. 1975. Range sites and soils in the United States. In: Hyder, D.N., ed. Arid shrublands–Proceedings of the 3rd workshop of the United States/Australia rangelands panel. 1973 March 26-April 5; Tucson, AZ. Denver, CO: Society for Range Management: 26-32.

    Shiflet, T.N. 1994. Rangeland cover types of the United States. Denver, CO: Society for Range Management. 152 p.

    Shock, C.C.; Feibert, E.B.; Saunders, L.D.; Parris, C.; Shaw, N.; Sampangi, R. 2011. Seed production of Great Basin native forbs- Subsurface drip irrigation (SDI) for stable, efficient native forb seed production using small amounts of supplemental irrigation water; weed control; and seeding practices. In: Shaw, N.L.; Pellant, M., eds. Great Basin Native Plant Selection and Increase Project: 2010 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 93-128.

    Shock, C.C.; Feibert, E.B.G.; Felix, E.B.; Shaw, N.L.; Kilkenny, F. 2015. Irrigation requirements for native perennial wildflower seed production in a semi-arid environment. In: Kilkenny, F; Halford, A.; Malcomb, A., eds. Great Basin Native Plant Project: 2014 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 101-122.

    Shock, C.C.; Feibert, E.B.G.; Rivera, A.; Saunders, L.D. 2018. Irrigation requirements for Lomatium seed production in a semi-arid environment. In: Kilkenny, F; Edwards, F.; Irwin, J.; Barga, S., eds. Great Basin Native Plant Project: 2017 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 97-110.

    Shock, C.C.; Feibert, E.B.G.; Rivera, A.; Saunders, L.D.; Shaw, N.; Kilkenny, F.F. 2016. Irrigation requirements for seed production of five Lomatium species in a semiarid environment. HortScience. 51(10): 1270-1277.

    Simmons, S.A.; Rickard, W.H. 2002. Plant succession at the edges of two abandoned cultivated fields on the Arid Lands Ecology Reserve. Northwest Science. 76(1): 85-89.

    Skinner, D. 2007. Propagation protocol for production of container (plugs) Lomatium triternatum (Pursh.) J.M. Coult. & Rose plants 10 cu in. Native Plant Network. U.S. Department of Agriculture, Forest Service, National Center for Reforestation, Nurseries, and Genetic Resources. http://npn.rngr.net/propagation/protocols [Accessed 2020 July 1].

    Skinner, D.M.; Warnick, P.; French, B.; Fauci, M. 2005. Characteristics and uses of native Palouse forbs in landscaping. Available from http://www.palouseprairie.org.

    Soltis, P.S.; Soltis, D.E.; Norvell, T.L. 1997. Genetic diversity in rare and widespread species of Lomatium (Apiaceae). 44(1): 59-73.

    Sosa, E.M.; Del Polito, L.; Stevens, M.; Mansfield, D.H.; Smith, J.F. 2015. Speciation in western North America: Lomatium as an example of diversification and convergent evolution. Boise State University, Poster. Available: https://scholarworks.boisestate.edu/cgi/viewcontent.cgi?article=1005&context=icur [Accessed 2020 July 1].

    St. John, L.; Ogle, D. 2011. Cooperative work between the Great Basin Native Plant Selection and Increase Project and the Aberdeen Plant Materials Center. In: Shaw, N.; Pellant, M., eds. Great Basin Native Plant Selection and Increase Project: 2010 Progress Report. Boise, ID: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 69-76.

    Stevens, R.; Monsen, S.B. 2004. Forbs for seeding range and wildlife habitats. In: Monsen, S.B.; Stevens, R.; Shaw, N.L., comps. Restoring western ranges and wildlands, vol. 2. Gen. Tech. Rep. RMRS-GTR-136-vol-2. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 425-466.

    Stuth, J.W.; Winward, A.H. 1977. Livestock-deer relations in the lodgepole pine-pumice region of central Oregon. Journal of Range Management. 30(2): 110-116.

    Taylor, R.J. 1992. Sagebrush country: A wildflower sanctuary. Missoula, MT: Mountain Press Publishing Company. 211 p.

    Thompson, J.N. 1985. Postdispersal seed predation in Lomatium spp. (Umbelliferae): Variation among individuals and species. Ecology. 66(5): 1608-1616.

    Tilley, D.; Bair, C. 2011. The Jet Harvester: A shop tool for harvesting forb and shrub seeds. Native Plants Journal. 12(2): 123-127.

    Tilley, D.; St. John, L.; Ogle, D.; Shaw, N.; Cane, J. 2010. Plant guide: Nineleaf biscuitroot (Lomatium triternatum [Pursh] Coulter & Rose). Aberdeen, ID: U.S. Department of Agriculture, Natural Resources Conservation Service. 4 p.

    Tilley, D.; Taliga, C.; Burns, C.; St. John, L. 2013. Plant materials for pollinators and other beneficial insects in eastern Utah and western Colorado. Tech. Note 2C. Boise, ID: U.S. Department of Agriculture, Natural Resources Conservation Service. 54 p.

    Tisdale, E.W. 1979. A preliminary classification of Snake River Canyon grasslands in Idaho. Stn. Note 32. Moscow, ID: University of Idaho, Forest, Wildlife and Range Experiment Station. 8 p.

    Tisdale, E.W.; Hironaka, M.; Fosberg, M.A. 1965. An area of pristine vegetation in Craters of the Moon National Monument, Idaho. Ecology. 46(3): 349-352.

    Turner, N.J.; Bouchard, R.; Kennedy, D.I. 1980. Ethnobotany of the Okanagan-Colville Indians of British Columbia and Washington. Victoria, BC: British Columbia Provincial Museum. 179 p.

    USDA Forest Service [USDA FS] 1937. Range plant handbook. Washington, DC: U.S. Department of Agriculture, Forest Service. 816 p.

    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, 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, 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]. 2019. 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: U.S. Department of the Interior, Bureau of Land Management, Plant Conservation and Restoration Program. 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 and Restoration Program.

    USDI Environmental Protection Agency [USDI EPA]. 2017. Ecoregions. Washington, DC: U.S. Department of the Interior, 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.

    Van Allen Murphey, E. 1990. Indian uses of plants. Glenwood, IL. Meyerbooks. 81 p.

    Vankus, V. 2006. Association of official seed certifying agencies (AOSCA). https://www.aosca.org/.

    Walker, S.C.; Shaw, N.L. 2005. Current and potential use of broadleaf herbs for reestablishing native communities. In: Shaw, N.L.; Pellant, M.; Monsen, S.B., comps. Sage-grouse habitat restoration symposium proceedings; 2001 June 4-7; Boise, ID. RMRS-P-38. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station: 56-61.

    Welsh, S.L.; Atwood, N.D.; Goodrich, S.; Higgins, L.C., eds. 2015. A Utah Flora. Fifth Edition, revised. Provo, UT: Brigham Young University. 990 p.

    Whittaker, R.H. 1960. Vegetation of the Siskiyou Mountains, Oregon and California. Ecological Monographs. 30(3): 279-338.

    Wiese, J.L.; Meadow, J.F.; Lapp, J.A. 2012. Seed weights for northern Rocky Mountain native plants with an emphasis on Glacier National Park. Native Plants Journal. 13(1): 39-49.

    Wilson, B.L. 2019. Key to Apiaceae of Idaho and Montana. Corvallis, OR: Carex Working Group. 35 p.

    Young, S.A.; Schrumpf, B.; Amberson, E. 2003. The Association of Official Seed Certifying Agencies (AOSCA) native plant connection. Moline, IL: AOSCA. 9 p. Available: https://seedcert.oregonstate.edu/sites/seedcert.oregonstate.edu/files/pdfs/aoscanativeplantbrochure.pdf

    How to Cite

    Gucker, Corey L.; Shaw, Nancy L. 2020. Nineleaf biscuitroot (Lomatium triternatum [Pursh] J.M. Coult. & Rose). In: Gucker, C.L.; Shaw, N.L., eds. Western forbs: Biology, ecology, and use in restoration. Reno, NV: Great Basin Fire Science Exchange. Online: https://westernforbs.org/species/nineleaf-biscuitroot-lomatium-triternatum/