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Sorghum halepense

• INTRODUCTORY
• DISTRIBUTION AND OCCURRENCE
• BOTANICAL AND ECOLOGICAL CHARACTERISTICS
• FIRE ECOLOGY
• FIRE EFFECTS
• MANAGEMENT CONSIDERATIONS
• REFERENCES

INTRODUCTORY

AUTHORSHIP AND CITATION FEIS ABBREVIATION SYNONYMS NRCS PLANT CODE COMMON NAMES TAXONOMY LIFE FORM FEDERAL LEGAL STATUS OTHER STATUS
	Craig Thornsen. © 2001, California Dept. of Food and Agriculture

DISTRIBUTION AND OCCURRENCE

• GENERAL DISTRIBUTION
• ECOSYSTEMS
• STATES/PROVINCES
• BLM PHYSIOGRAPHIC REGIONS
• KUCHLER PLANT ASSOCIATIONS
• SAF COVER TYPES
• SRM (RANGELAND) COVER TYPES
• HABITAT TYPES AND PLANT COMMUNITIES

In the United States, Johnsongrass was introduced in South Carolina from Turkey around 1830. William Johnson, whom the plant is named after, established Johnsongrass along the Alabama River in the 1840s as a forage species, and Johnsongrass spread rapidly across the South [14,149,170,182]. Johnsongrass is now widely escaped from cultivation in much of the United States. It is most invasive in the Southeast, although it is widespread in central California and New Mexico [122,128,206]. Johnsongrass is not persistent in the Pacific Northwest, upper northern Great Plains, extreme northern portions of the Great Lake states, the Northeast [70,110,132,194], or in Arizona, Colorado, and Utah [111,200,201]. Plants database provides a state distributional map of Johnsongrass.

Johnsongrass occurrence is not well documented for all plant communities where it may occur. The following classification lists are not restrictive, but include plant communities where Johnsongrass is a documented species.

Riparian and wetland associates: A vegetation survey on the lower Rio Grande of Texas found Johnsongrass was the most common herbaceous cover (34%) in Fremont cottonwood-Goodding willow (Populus fremontii-Salix gooddingii) communities, followed by docks (Rumex spp.) (32% cover). Saltcedar (Tamarix chinensis ) and seepwillow (Baccharis spp.) were most common in the shrub layer [56]. On the Tensas River National Wildlife Refuge, Louisiana, Johnsongrass is waa dominant herbaceous species (8% cover) in winged elm-American elm-cedar elm-green ash (Ulmus alata-U. americana-U. crassifolia-Fraxinus pennsylvanica) woodlands. Other dominant herbs included purple loosestrife (Lythrum salicaria, 21% cover), trumpet creeper (Campsis radicans, 13% cover), and bushy bluestem (Andropogon glomeratus, 7% cover) [123].

Old fields: On old bottomland fields of the Mississippi and Yazoo rivers, Mississippi, Johnsongrass cover was greatest on silty-clay loams. Associated overstory species were Texas red oak (Quercus texicana), Shumard oak (Q. shumardii), and cherrybark oak (Quercus  pagoda). Canada goldenrod (Solidago canadensis) was a common herbaceous associate [3]. In Oklahoma, Johnsongrass dominates on bottomlands and old fields, associating with ragweed (Ambrosia trifida), Canadian horseweed (Conyza canadensis), and smooth sumac (Rhus glabra). Hoagland [95] provides a description of Johnsongrass-dominated old-field communities in Oklahoma.

BOTANICAL AND ECOLOGICAL CHARACTERISTICS

GENERAL BOTANICAL CHARACTERISTICS RAUNKIAER LIFE FORM REGENERATION PROCESSES SITE CHARACTERISTICS SUCCESSIONAL STATUS SEASONAL DEVELOPMENT
	Entire Johnsongrass plant (left), and Johnsongrass rhizomes and culm shoots (right). Photos by Allan Kates, Virginia Polytechnic Institute (in [84]).

GENERAL BOTANICAL CHARACTERISTICS:
The following description of Johnsongrass provides characteristics that may be relevant to fire ecology, and is not meant for identification. Keys for identification are available (e.g., [72,93,94,96,111,149,176,194]).

Morphology: Johnsongrass is a nonnative, warm-season perennial [49,71,122,184,201]. It is usually rhizomatous, but is a highly variable species with many ecotypes [96]. It may grow as an annual in hot, arid climates and at the northern limits of its range [61,197,198]. Johnsongrass rhizomes form a dense, tangled, tough sod [204]. Rhizomes serve as carbohydrate-storing and regeneration organs [6]. Most rhizomes occur in the top 7.9 inches (20 cm) of soil, although rhizomes in soft, deep soil may extend deeper [42,96]. Rhizomes vary in size from a few inches to several feet in length, and in thickness from 0.25 to 0.75 inch (6.4-19 mm) [86]. Leaves and aboveground stems (culms) are coarse [149]. Culms are 1.6 to 4.9 feet (0.5-1.5 m) tall. Total plant height may reach 12 feet (3.7 m) during flowering [5]. The inflorescence is a 4- to 24-inch (10-60 cm) open panicle.  Spikelets of Sorghum species are paired: 1 is sessile and perfect; the other spikelet is pedicelled and staminate. Spikelets are 4 to 7 mm in length. There are about 35 to 350 spikelets per panicle, depending upon ecotype. Lemmas are cilate; they may be awnless or have short (1-15 mm), sometimes twisted awns that aid in seed dispersal [49,70,93,122,184,194,196,201]. Seeds are about 2 mm long [149].

Physiology: Several physiological characteristics of Johnsongrass aid in its spread. Mature Johnsongrass plants are moderately drought resistant [6] and salt tolerant [207]. Johnsongrass produces toxins (see Toxicity) that may be allelopathic [96,135,196].

Asexual regeneration: Once a population of Johnsongrass is established, most population growth is from asexual regeneration by rhizomes [96]. Throughout most of their North American range, Johnsongrass populations are strongly rhizomatous [49,71,122,184,201]. Some Johnsongrass populations are weakly rhizomatous or nonrhizomatous, especially at the species' distributional limits [61,197,198]. Rhizome expression in Johnsongrass is apparently controlled by multiple, dominant genes, resulting in variable degrees of rhizome development in both Johnsongrass and its hybrids [209]. Extreme temperatures also inhibit Johnsongrass's ability to produce rhizomes [85,175].

Rhizome development: Johnsongrass plants begin growing rhizomes in the seedling stage. Primary rhizomes are initiated at the 5-leaf stage, when plants are about a foot tall. Rhizome growth continues slowly until the 10-leaf stage, then accelerates greatly. Rhizomes are well developed by 6 to 7 weeks [6,101,125]. In a greenhouse experiment, Anderson and others [6] noted extensive rhizome development on 4.5-month-old plants, with over 5,200 rhizome nodes/plant.

In older plants, last-year or primary rhizomes produce new, secondary rhizomes in spring. Secondary rhizomes in turn produce tertiary rhizomes. Secondary and tertiary rhizome growth slows or stops during flowering, then resumes with seedhead development [126]. Rhizome production peaks at seed ripening [78,97], when a single plant may produce 200 to 300 feet (60-90 m) of rhizomes [127]. Secondary and tertiary rhizomes continue growth and carbohydrate accumulation until late fall, then go dormant over winter. Primary rhizomes die each fall. In spring, secondary and tertiary rhizomes become current-year primary rhizomes [4,42,80,86,96,101].

Rhizome sprouting: Small or broken rhizomes, especially secondary rhizomes, can form new plants [6,9,42,173]. Plows spread Johnsongrass by breaking up, dispersing, and replanting rhizomes [3] (see Physical/mechanical control). Small rhizomes are more likely to sprout when shallowly buried, while large rhizomes are more likely to sprout when deeply buried. In the greenhouse, 3-inch (7.6-cm) rhizome sections sprouted best when planted less than 3 inches deep. Longer, 6-inch (15.2-cm) sections sprouted best when buried deeper than 3 inches ([39] and references therein). In a Mississippi field experiment, McWhorter [125,126] found that with shallow burial (<2 inches (6 cm)), short rhizomes (<3 inches (7.6 cm)) produced more sprouts than long rhizomes (6 inches (15.2 cm)). The opposite trend occurred when rhizomes were planted deeper than 2.4 inches (6 cm). Rhizomes usually grow to a depth of 10 to 20 inches (25-50 cm) [127]. Loose, sandy or loamy soils generally allow for best rhizome expansion [126]. Clay tends to inhibit rhizome expansion [86]; however, rhizomes may penetrate several feet down cracks in clay soil [126]. Deeply buried rhizomes that do not sprout do not survive more than a year [4,80,96].

Rhizomes are somewhat drought-resistant, remaining viable after drying to 40% of initial harvest weight [6]. They are sensitive to extreme temperatures. In northern climates, rhizomes must be deeply buried in order to overwinter. In an Illinois field experiment, Johnsongrass rhizomes did not survive winter temperatures less than 1.4 °F (-17 °C) unless buried 7.9 inches (20 cm) or more below ground [175]. In southern Ontario, rhizomes must be 10 inches (25 cm) or more inches below ground to overwinter ([85] and references therein).

Sexual regeneration: Although growth of established populations is primarily through rhizomes, Johnsongrass establishes new populations through seed spread [96].

Breeding system: Sorghum species are mostly self-crossed, although some outcrossing occurs ([196] and references therein).

Pollination: Johnsongrass is primarily self-pollinated [196]. Some pollination is effected by wind, especially when plants are <425 feet (130 m) apart [196,202].

Seed production: Johnsongrass is a short-day plant, requiring 8 to 16 hours of daylight to flower [66,135]. It is a good seed producer under favorable growing conditions. A single plant may produce 80,000 or more seeds in 1 growing season [3,84]. Two seed crops may be produced under good conditions. In agricultural fields in Argentina, Johnsongrass produced a large seed crop in early summer (Jan-Feb.; 60% of total seed production for the year) and a smaller seed crop in late summer-early fall (mid-March-early April; 40% of annual seed production) [66]. Johnsongrass seed production is estimated at 90 gallons/acre (855 l/ha) on good sites in the South. Field trials in Mississippi showed mean seed production of 84 g/plant and 28,000 seeds/plant [127,196]. Resources are allocated to rhizomes at the expense of seeds under poor growing conditions [22,23].

Greenhouse trials using Johnsongrass seed from the Northeast showed populations that grow and reproduce as annuals have faster growth rates, more rapid development, more and larger seeds, and fewer rhizomes compared to populations that sprout from overwintered rhizomes [198].

Seed dispersal: Wind, water, machinery, and animals disperse Johnsongrass seeds [3,65,84,85,182]. Spikelets are readily deciduous [201] and usually disperse as a unit beneath the parent plant [3,65,70,72]. Strong winds disperse seeds longer distances. In Argentina, 28- to 31-mile/hr (45- to 50-km/hr) winds that occurred during May thunderstorms carried Johnsongrass seeds 2,950 to 3,300 feet (900-1,000 m) from parent plants [65]. Water has dispersed seeds along many waterways of the United States [3,84,182]. Farming equipment also spreads seeds [65,84]. Viable Johnsongrass seed is a common contaminant in hay, harvested crops, and commercial seed [3,142]. Johnsongrass seed retains viability after passing through the digestive tracts of livestock [9,84,126]. The relative importance of agents that disperse Johnsongrass seed is unclear [128].

Seed banking: Johnsongrass builds up a soil seed bank [189]. The seeds are dormant and may remain viable for several years, although most soil-stored seeds germinate in their 1st or 2nd year [3,96,135]. In Mississippi, 1st-year stratified seed showed 82% viability in the field. After burial in the field for 2.5 years, the same seed lot showed 62% viability [53]. In California, 5-year-old buried seed showed >50% viability, but viability dropped to 2% by age 6 [120].

Germination: Johnsongrass has 2 mechanisms of dormancy: mechanical dormancy imposed by the seed hull and seedcoat, which requires weathering or scarification to break; and chemical dormancy, which requires oxygen to break [100]. Diurnal fluctuations in temperature, afterripening, or both are needed to overcome both types of dormancy [26,26,64,99,181]. Seed from water-stressed plants is generally less dormant than seed from amply watered plants [23]. Benech and others [25] present a model predicting loss of seed dormancy and consequent seedling emergence based on soil temperature.

Light improves germination rate with warm temperatures (>93 °F (34 °C)) and inhibits germination with cold temperatures (<72 °F (22  °C) [100]. In the greenhouse, Taylorson and McWhorter [181] found a 63% increase in germination rate for Johnsongrass seed exposed to light vs. seeds kept in the dark. Deeply buried seed remains dormant for at least 7 years [84] but does not germinate [27,64,99]. Soil upheaval such as cultivation, which brings seed closer to the soil surface, usually increases germination rates [64,99]. In the greenhouse, best germination (60-75%) occurred with surface-scattered to shallowly buried (0-1.6 inches (0-4 cm)) seed. Less than 5% germination occurred with seed buried >3 inches (8 cm) below the soil surface [27]. Litter cover or shallow burial may aid germination in the field. Prostko and others [147] present a model to predict Johnsongrass seedling emergence based upon temperature and seed burial depth.

Seedling establishment/growth: Best establishment occurs on open, disturbed sites. Seed dispersed away from parent plants may show better establishment compared to seed falling beneath the parent. In an old field in Argentina, most Johnsongrass seed fell near parent plants on undisturbed plots. Only 1% of seed beneath a parent plant established. On tilled plots mowed every 1 to 2 weeks by a corn (Zea mays) harvester, seed was carried 3 to 82 feet (1-25 m) from parent plants. Recruitment of tilled seed neared 100% [65]. On favorable sites, plants may produce 80 or more culms in their 1st year [3].

In areas where Johnsongrass grows as a facultative annual, it shows variable ability to regenerate from seed. Johnsongrass annuals in rural-interface wildlands of southern Arizona rarely reproduce either from on-site seed or from rhizomes. Seed dispersed from adjacent agricultural lands provide continual sources of seed [61]. However, some annual populations in the northern portion of Johnsongrass's range successfully reproduce from seed. In southern Ontario, northern Ohio, and northern New York, annual populations have larger leaves, inflorescences, and seeds compared to perennial Johnsongrass populations [198].

Johnsongrass seedlings may show faster 1st-year growth than plants started from rhizome fragments. On the Mississippi Delta near Stoneville, Mississippi, Johnsongrass started from seed showed greater biomass and more rapid height gain than plants started from rhizome pieces. At flowering, seed plants were producing 0.75 to 3 feet (0.23-0.9 m) of new rhizome growth per day. Plant growth patterns were as follows (data are means) [125]:

Moisture regime: Although Johnsongrass occurs in wet to dry habitats in its native range in southern Europe [184], it is generally restricted to wet or mesic sites in the United States [72,93,201]. It is most common in warm, humid southern climates that receive ample summer rainfall. Johnsongrass is a facultative wetland species, frequently occurring on floodplains [18,19]. Johnsongrass patches are often extensive along canals and irrigation ditches [96]. In arid sites such as Organ Pipe Cactus National Monument, Arizona, Johnsongrass establishes mostly in wet locations including washes, drainages, and stream edges [61,177].

Elevational ranges of Johnsongrass in several states are:

Old fields: Johnsongrass is particularly common on old bottomland fields in the South [3,17,18]. Unlike most crop weeds, which tend to decrease in the absence of irrigation and fertilization, Johnsongrass tends to persist on abandoned fields. For example, in Georgia it was prevalent in 1-, 4-, and 8-year-old fertilized crop fields, but was also prevalent in 8-year-old fallow fields [139].

Shade tolerance: Johnsongrass requires open sites and does not persist under closed canopies [77,190]. In a honey mesquite (Prosopis glandulosa) Texas savanna, Johnsongrass associated with Texas wintergrass (Nassella lecotricha) in open areas but was not found under honey mesquite or other trees [190]. In Argentina grasslands, canopy removal increased Johnsongrass germination and establishment compared to closed-canopy sites [25].

Johnsongrass produces seed about 2 months after initiation of spring rhizome expansion [151]. Even 1st-year Johnsongrass plants are capable of quick flowering and seed set: Plants in Mississippi initiated flowers 46 days after seedling emergence [125]. Bridges and Chandler [35] present a model for predicting Johnsongrass flowering under fluctuating field temperatures, with nitrogen fertilization, and with irrigation. Holshouser and Chandler [97,98] provide temperature-dependent models for predicting flowering, germination, rhizome bud break, and rhizome sprouting under field conditions.

The following table shows phenological development of Johnsongrass in various states and regions:

FIRE ECOLOGY

• FIRE ECOLOGY OR ADAPTATIONS
• POSTFIRE REGENERATION STRATEGY

Little documentation is available on postfire regeneration of Johnsongrass from seed. As a seed banking species that produces abundant, readily dispersed seed (see Sexual regeneration) that establishes well in open, disturbed sites (see Seedling establishment/growth), it is likely that Johnsongrass is capable of postfire seedling establishment from both on- and off-site sources. Germination of Johnsongrass seed was not affected by exposure to temperatures of 200 °F (90 °C) and 400 °F (200 °C) for 120 seconds. However, germination was dramatically reduced following exposure to 660 °F (350 °C) and was eliminated after exposure to 800 °F (430 °C) and higher [131]. See [131]. See Immediate Fire Effect on Plant for details of this study. More information is needed on postfire establishment potential of Johnsongrass.

Fuels: Johnsongrass litter reportedly remains on the ground all winter [104]. Johnson [107] provides a simple technique for estimating ratios of live:dead plant materials in Johnsongrass.

Fire regimes: As of this writing (2004), there was no published information on how North American fire regimes affect Johnsongrass. In riparian and other areas where Johnsongrass is highly productive, Johnsongrass may promote fire spread by increasing fine fuel loads above historical levels. Studies are needed on the fire ecology of Johnsongrass in North American.

The following table provides fire return intervals for plant communities and ecosystems where Johnsongrass may be important. Find further fire regime information for the plant communities in which this species may occur by entering the species name in the FEIS home page under "Find Fire Regimes".

FIRE EFFECTS

• IMMEDIATE FIRE EFFECT ON PLANT
• DISCUSSION AND QUALIFICATION OF FIRE EFFECT
• PLANT RESPONSE TO FIRE
• DISCUSSION AND QUALIFICATION OF PLANT RESPONSE
• FIRE MANAGEMENT CONSIDERATIONS

Fire scarification appears to have no effect on rate of seed germination, and very high temperatures kill Johnsongrass seed. In the laboratory, there were no significant differences in rates of germination between unheated Johnsongrass seed and seed heated to 200 °F (90 °C) and 400 °F (200 °C) for 120 seconds; mean germination rate was 52%. Mean germination rate dropped to 17% for seed exposed to 660 °F (350 °C) and to 0% for seed exposed to 800 °F (430 °C) and 1000 °F (538 °C) for 120 seconds [131].

Published information on postfire seedling establishment of Johnsongrass is lacking. Studies are needed on the ability of Johnsongrass to establish from seed in postfire environments.

Spot burning with a gas torch was once a widely practiced method of controlling patches of Johnsongrass. It is rarely used today due to cost restraints [127], but may be useful in riparian or other areas where spot control is needed. In Arizona, a Johnsongrass infestation along a canal bank was controlled by torching plants with a propane burner. Eleven burnings at 2-week intervals controlled top growth. No regrowth of Johnsongrass occurred the following season [86].

MANAGEMENT CONSIDERATIONS

• IMPORTANCE TO LIVESTOCK AND WILDLIFE
• OTHER USES
• IMPACTS AND CONTROL

Palatability/nutritional value: Johnsongrass is moderately palatable and nutritious [47,49]. Deer make light to moderate use of Johnsongrass [114,163], grazing all aboveground portions of the plant [47]. Rodents also graze Johnsongrass. In honey mesquite (Prosopis glandulosa var. glandulosa) plains of Texas, Heerman's kangaroo rat and Great Basin pocket mouse used Johnsongrass frequently (6.7% volume, 61% frequency; and 5.9% volume, 36% frequency, respectively) [2]. Quail, geese, and wild turkey consume Johnsongrass seeds [31,47].

Although intolerant of heavy grazing, Johnsongrass is a good pasture grass and makes fair-quality hay when cut in the boot stage [170,191]. Livestock make moderate to good use of fresh Johnsongrass [87,91]. In a comparison of Texas range grasses, Johnsongrass showed greatest in-vitro digestibility (45%-69%, depending on the digestion media) of 5 grasses tested [116]. Dairy cattle in Alabama showed good weight gain and milk production on Johnsongrass pasture [87].

Nutritional content: In a western Texas study, Johnsongrass had highest summer crude protein content (13.2%) of 9 rangeland grasses. Spring, fall, and winter crude protein values were 6.62%, 8.06%, and 3.81%, respectively [37]. In a greenhouse study comparing relative mineral content of 15 grass species, Johnsongrass scored significantly higher in phosphorus content than other grass species (P=0.1). Cobalt, manganese, and copper contents were moderate compared to other grasses [29]. Energy value of Johnsongrass grown in Texas was 3,900 kcal/g [136]; in India, seasonal fluctuation in energy value varied from 3,684 kcal/g in October to 4,578 kcal/g in April [167]. Nutritional content of fresh Johnsongrass in eastern Texas was [62]:

Seasonal changes in forage quality of Johnsongrass on the Edwards Plateau of Texas were [103]:

When harvested at its peak, Johnsongrass makes fair-quality hay, similar to timothy (Phleum pratense) hay in nutrient content [146]. In a 1928 study in an Alabama coal mine, draft horses and mules were fed oat (Avena sativa) grain and either Johnsongrass or timothy hay for 3 months. The equines maintained their weight on both diets under "moderate" workloads. The animals lost weight on both diets under "heavy" workloads, but lost less weight on timothy hay and grain compared to Johnsongrass hay and grain (mean losses of 10.71 and 21.78 lbs., respectively). Digestible nutrient means were [75]:

Toxicity: Johnsongrass is generally a good forage grass [31,47,87,91]. However, at certain developmental stages or under some adverse environmental conditions, Johnsongrass may form cyanogenetic glycosides that can poison livestock. Phenologically, Johnsongrass is most toxic when leaves and culms are actively growing. Seedlings and sprouts generally have higher levels of glycosides than plants that have reached the flowering stage. Secondary growth, produced after mature plants are mowed or heavily grazed, can also have high levels of glycosides. Environmentally, Johnsongrass is most toxic after drought, extreme heat, frost, or when plants are wet with dew or light rain. Glycoside levels can vary considerably among Johnsongrass populations. Ruminants, especially cattle, are more susceptible to glycoside poisoning than monogastric herbivores like horses  [157,173]. As well as fresh plants, hay cut when Johnsongrass is young or experiencing adverse environmental conditions such as drought can also be toxic [81,133,170,170]. Livestock poisoning can be prevented by waiting until new growth is 15 to 18 inches tall (38-46 cm) tall after drought, or deferring grazing until plants have dried after frost [128].

Johnsongrass may sequester selenium or other elements that are toxic at high doses when growing in soils with high concentrations of toxic elements. In the Dead Sea area of Jordan, for example, selenium concentrations in Johnsongrass samples were high enough to poison livestock [1].

Prolonged consumption of fresh Johnsongrass can cause nitrate poisoning in ungulates [173]. Most livestock can graze Johnsongrass safely when plants are at least 18 inches (46 cm) tall [170].

Cover value: No information is available on this topic.

Little is documented on Johnsongrass's impact in wildlands, and further research is needed on how Johnsongrass affects wildland habitats. Generalizations about Johnsongrass must always be qualified because of numerous ecotypes [125]. Typically, Johnsongrass is a good competitor for nutrients [96,196], space [103], and water [166]. It can outcompete associated species for water by extracting water from lower soil profiles (12 inches (30 cm) or more below ground) [105]. Johnsongrass may also negatively impact plant community composition through its reputed allelopathy [96,135,196]. Cyanogenetic glycosides and other toxins in Johnsongrass may inhibit germination and growth of associated plant species [96,135,196].

On many sites in the United States, Johnsongrass is not invasive in undisturbed wildlands, although it may readily invade disturbed sites (Cox, cited in [135]). Johnsongrass is most invasive on moist sites in wet-temperate regions of the southeastern United States [71,72,135]. For example, Johnsongrass and Canada thistle (Cirsium arvense) were listed as the 2 most invasive and expensive to control weeds on the Eastern Neck National Wildlife Refuge, Maryland [44]. Johnsongrass interferes with conifer seedling establishment and growth on southern pinelands [50], and may interfere with cottonwood (Populus spp.) and willow (Salix spp.) establishment in riparian zones [171].

Johnsongrass is not invasive on most sites in the Southwest. At the turn of the last century, Johnsongrass was planted in southwestern arroyos and stream channels to stabilize soil [46]. It established on such wet and mesic sites, but failed to spread. Felger [61] reports Johnsongrass as only weakly invasive in Organ Pipe Cactus National Monument, Arizona, where the arid climate restricts Johnsongrass to roadsides and washes. Johnsongrass may grow as an annual, without spreading, in arid southwestern wildlands. Nearby agricultural lands are continual seed sources [61].

Control: Although considerable information is available on controlling Johnsongrass in agricultural settings (e.g., see [9,80,84,127,180]), information on controlling Johnsongrass in rangelands, natural areas, and other wildlands is lacking. The following information on Johnsongrass control is extracted primarily from agricultural literature but may be applied to some wildland settings, particularly old fields. Research is needed on controlling Johnsongrass in wildland settings [135].

Johnsongrass control involves several steps: 1) preventing seed from ripening and dispersing, 2) killing seedlings, 3) killing existing rhizomes, and 4) preventing growth of new rhizomes [9,80,84,127,180]. Control is most effective before plants have developed 5 leaves [101]. Detailed Johnsongrass control procedures and techniques are given in several publications [96,127,135].

Prevention:  The most efficient and effective method of managing invasive species such as Johnsongrass is to prevent their invasion and spread [164]. Preventing the establishment of nonnative invasive plants in wildlands is achieved by maintaining native communities and conducting aggressive surveying, monitoring, and any needed control measures several times each year. Monitoring efforts are best concentrated on the most disturbed areas in a site, particularly along potential pathways for Johnsongrass invasion: roadsides, waterways, and old fields. Large plant size makes monitoring Johnsongrass relatively easy in summer, and yearly summer monitoring helps managers assess the effectiveness of control programs. As of this writing (2004), monitoring programs for Johnsongrass were in their infancy. As potential contact sources, Newman [135] provides a list of managers who have started monitoring programs for Johnsongrass on Natural Areas. The Center for Invasive Plant Management provides an online guide to noxious weed prevention practices.

Integrated management: A combination of complementary control methods may be helpful for rapid and effective control of Johnsongrass. Integrated management includes not only killing the target plant, but establishing desirable species and discouraging nonnative, invasive species over the long term. Johnsongrass control is rarely successful with only 1 method of control [141], but a combination of control methods can be effective. For example, in a tallgrass restoration study on the Hear Wildlife Sanctuary, Texas, a combination of early fall glyphosate spraying followed by late fall tillage helped control nonnative grasses on a former Johnsongrass-Bermuda grass (Cynodon dactylon) pasture. Early fall spraying targeted Johnsongrass while it was still actively growing. After spraying, rhizomes brought to the soil surface by tilling 4 to 6 inches (10-15 cm) deep were killed by winter frost. Johnsongrass showed 4.2% cover and 50% frequency 3 years after treatments. Only trace amounts of Bermuda grass were present [172].

Fire: See Fire Management Considerations.

Biological: Biological control of Johnsongrass is problematic, as known control agents that kill Johnsongrass also kill crop grasses such as corn and sorghum [128,143,145]. As of this writing (2004), there are no biocontrol agents approved for Johnsongrass [183]. Several biological agents are being tested for possible use. A smut (Sphacelotheca holci) has helped control Johnsongrass in Louisiana croplands [130]. In Florida field trials, a mixture of native fungal pathogens controlled Johnsongrass and other weedy grasses in citrus (Citrus spp.) groves [43].

Heavy grazing over 2 or more years reduces Johnsongrass by depleting rhizome reserves [3,8,89]. Rhizome development is greatly reduced when plant height is kept below 12 to 15 inches (30.5-38 cm) [127]. Best control is offered when herbicide or winter plowing treatments follow grazing treatments [3]. For example, in an unpublished study at the Patagonia/Sonoita Creek Preserve, Arizona, cow and horse summer grazing reduced density of Johnsongrass. After 4 years of summer grazing, Johnsongrass stem density had decreased 75% compared to pretreatment levels. Plots were then sprayed in late spring with glyphosate. Posttreatment restoration plantings gave mixed results. One to two months after spraying, native bunchgrasses were transplanted onto the study sites. Broadleaf weeds invaded the study plots after Johnsongrass density was reduced by the grazing and herbicide treatments. After mowing treatments to control the broadleaved weeds, native bunchgrasses on some test plots were showing good growth. Other plots experienced Johnsongrass reinvasion and pocket gopher herbivory, to the detriment of native bunchgrasses. Preserve managers are continuing weed control treatments to promote the native bunchgrasses [183].

Geese are sometimes used for Johnsongrass control in croplands. Geese prefer young shoots, and do not graze Johnsongrass over about 7 inches (18 cm) in height [9,86].

Chemical: Herbicides may provide initial control of a new invasion or a severe infestation, but used alone, they are rarely a complete or long-term solution to invasive species management [40]. Herbicides are most effective on large infestations when incorporated into long-term management plans that include replacement of weeds with desirable species, careful land use management, and prevention of new infestations. Control with herbicides is temporary, as it does not change the conditions that allowed the invasion to occur in the first place (e.g., [211]). See The Nature Conservancy's Weed Control Methods Handbook for considerations on the use of herbicides in Natural Areas and detailed information on specific chemicals.

The most effective chemical control of Johnsongrass involves using systematic herbicides that translocate the active chemicals to rhizomes [127]. A single application of herbicide generally does not control large infestations, and follow-up measures are needed for long-term control [169]. Johnsongrass control can be obtained using glyphosate [7,21,104,127], phenoxy (e.g., 2,4-D, fluazifop), [108,115], or halogenated aliphatic (e.g., dalapon) herbicides [8,86,127]. Spot spraying with sodium chlorate [82,86] or dalapon has been effective for small infestations [153]. Spot control is not effective in the long term unless surrounding seed sources are also eliminated [104]. Experiments in agricultural fields in Argentina showed best control when the herbicide (dalapon) was applied when rhizome biomass was low. Ghersa and others [67] provide a model for predicting optimal spraying time based on minimum rhizome biomass. Although based on South American seasons, the model is easily adjustable for use in the northern hemisphere.

Postemergent herbicides are the most common method of Johnsongrass control in agricultural systems, and are probably the best herbicide choice for wildland settings as well, since postemergent herbicides cause less damage to nontarget species. In a Maryland old-field study, foliar application of postemergence herbicide (DPX-V9360) was more effective in late-growth stages (>5 leaves) than early-growth stages (<5 leaves) when rhizomes had not fully expanded [138]. Rosales-Robles and others [158] discuss the relative effectiveness of several postemergent herbicides as influenced by application rate and Johnsongrass growth stage. Application procedures for postemergent herbicides effective on Johnsongrass are given in these publications: [20,51,119,208].

Ecotypes may show differential response to herbicides [128]. Populations in Kentucky and Mississippi show genetic resistance to fluazifop and other phenoxy herbicides [15,137,168]. Virginia populations have resistance to enzyme acetyl-coenzyme A carboxylase inhibitors [33]. In Greece, some populations show resistance to glyphosate [113].

Herbicide treatments greatly decreased Johnsongrass cover in an Illinois bottomland old field. Restoration treatments included tillage, pre- or postemergent herbicide applications (sulfometuron or glyphosate, respectively), and green ash (Fraxinus pennsylvanica) plantings. Tillage had no significant impact on Johnsongrass cover. Mean Johnsongrass cover (%) was significantly lower after the 1st postspray year [76]:

Cultural: Little information is available on cultural methods of control for Johnsongrass. An Arizona study using integrated pest management, including native bunchgrass plantings, showed some success in controlling Johnsongrass (see grazing in the Biological control section above). Additional studies incorporating cultural control of Johnsongrass are needed.

Physical/mechanical: Johnsongrass can be controlled by tilling, mowing, and flooding [6,127,169]. Individual small plants or small clumps may be controlled by hand-pulling or solarization [13,54,169].

A consistent tillage program may provide effective control [6,42,80,125]. Tilling is not practical on most wildlands due to damage to desirable native plant species, uneven terrain, erosion, and cost constraints [104]. Tilling can be used on some sites such as bottomlands and old fields. Shallow plowing helps control Johnsongrass by breaking up rhizome systems, exposing rhizomes to the sun or killing frosts, and depleting carbohydrate reserves [6,42,80,125]. Optimal plow depth is 8 to 12 inches (20-30 cm). Several treatments are needed in hot climates [86,101]. Killing sprouts early, before they form 5 leaves and start developing new rhizomes, gives best control [101]. First plowing is in spring (May), followed by similar plowings every 3 weeks (in rainy weather) to 6 weeks (in dry weather). Plant heights of 12 or more inches (30 cm) are recommended before plowing again [9]. In cold climates, Johnsongrass is plowed in late October to expose rhizomes to frost [19]. An exposure of 24 or more hours to temperatures below 25 °F (- 4 °C) kills rhizomes [80,102,125,126]. A single plowing, or long intervals between plowings (>4 years), is generally not effective because it stimulates growth [104,170], buries and protects rhizomes [42], and exposes deeply buried seeds to upper soil levels where they may germinate [64].

Because rhizomes may extend more than 20 inches (51 cm) below ground, cultivation alone may fail to kill Johnsongrass rhizomes [42]. After plowing, close grazing or mowing (so that the grass stays <12-15 inches (30-38 cm) tall) helps further reduce Johnsongrass cover [86].

Even on old fields, tilling is a major soil disturbance that provides a favorable seedbed for pioneer species. Unless further rehabilitation efforts that include planting native herbaceous species are taken, it is likely that tilled fields will succeed to other invasive nonnatives.

Repeated, close mowing has the same inhibitory effect on growth as grazing [104,169]. In Mississippi, mowing seedlings 13 days after emergence killed them [125]. In an Alabama field experiment, multiple cuttings, starting when plants were 1 foot (0.3 m) high, slowed Johnsongrass rhizome development. At the end of the growing season, plots cut 8 times averaged 15 dry-weight ounces (431 g) of Johnsongrass top-growth and 0.3 dry-weight ounces (10 g) of rhizomes. Plants cut only twice had 67 ounces (1,909 g) top-growth and 26 ounces (739 g) of rhizomes. Plots were 4 × 5 feet²[179].

Flooding for 3 to 6 weeks in early spring, before rhizomes sprout, can effectively control Johnsongrass. Replacing open irrigation ditches with culverts or pipes helps prevent reinfestation [127].

Hand-pulling Johnsongrass usually leaves rhizome pieces behind in the soil, stimulating sprouting. It is not an effective control method unless all rhizomes are removed or new sprouts are controlled [104,169]. Best results are obtained in early spring when soil in moist and rhizomes are least likely to break [169].

Repeated solarization treatments (using a clear polyethylene tarp to trap solar heat in the soil) can control small Johnsongrass infestations [13].

Seeds: Solarization of moist soil at 140 °F to 150 °F (60-70 °C) for 7 days kills most Johnsongrass seeds. Solarization of dry soil does not kill Johnsongrass seed [54]. In Davis, California, soil watered and solarized for 9-12 weeks supported no Johnsongrass. Untreated control plots showed 58% Johnsongrass cover [55]. For established plants, 30 days of solarization kills most Johnsongrass. Remaining plants have grown rhizomes through and above the landscape fabric, but rhizomes above the landscape fabric were easily removed by hand-pulling [121].

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