Le Réseau BIO

Plate-forme de réseautage pour les producteurs, transformateurs et commerçants d'aliments biologiques du Québec
Le Réseau BioUn site réalisé grâce à un partenariat
CETAB+ | Centre d'expertise et de transfert en agriculture biologique et de proximitéMinistère de l'Agriculture, des Pêcheries et de l'Alimentation du Québec
Bienvenue sur le Réseau BIO, une plate-forme de réseautage pour les producteurs, transformateurs et commerçants d'aliments biologiques et intervenants en agriculture biologique au Québec.

Modifier eXtension Articles,News,Faqs,Events- organic production (anglais)

S'abonner à flux Modifier eXtension Articles,News,Faqs,Events- organic production (anglais)
Mis à jour : il y a 1 heure 45 min

Palmer Amaranth (Amaranthus palmeri)

mer, 2012/09/12 - 16:56

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Introduction

Palmer amaranth, also known as Palmer pigweed, is an extremely aggressive, fast-growing species that has become a serious weed problem in vegetable and row crops in the southern half of the United States in recent years. Native to the Sonoran Desert and the lower Rio Grande Valley (Ehleringer, 1983; Keely, 1987), Palmer amaranth readily invades croplands in hot climates. It became a major agricultural weed in the southern Great Plains by the late 1990s (Horak, 1997), and now infests at least 750,000 acres of cotton and other row crops in Arkansas, (Fugate, 2009) and over one million acres in Georgia (Langcuster, 2008). In addition, it has been cited as a major troublesome weed in vegetable production in North and South Carolina (Webster, 2006). Over the past 10 years, numerous reports have been published on Palmer amaranth documenting severe crop losses, and resistance to glyphosate and other herbicides (Culpepper et al., 2006; Horak and Peterson, 1995; Jha et al., 2008a, b).

Palmer amaranth is a tall, erect, branching summer annual, commonly reaching heights of 6–8 feet, and occasionally 10 feet or more. Stems and foliage are mostly smooth and lacking hairs (glabrous). Leaves have fairly long petioles and are arranged symmetrically around the stem, giving the plant a distinctly pointsettia-like appearance when viewed from above (Fig. 1). Leaf blades are elliptical to diamond-shaped with pointed tips, and measure 0.6–3 inches long by 0.4–1.5 inches wide.

Palmer amaranth
Figure 1. (a) Palmer amaranth in vegetative growth stage, showing pointsettia-like growth habit. (b) Palmer amaranth at early head emergence, showing smooth, hairless foliage and stems. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Male and female flowers are borne on separate plants (dioecious), and the small (<0.25 inch) flowers are clustered tightly in linear or sparingly branched terminal spikes up to 18 inches long (Fig. 2). The perianth (whorl of petal-like structures) around each female flower bears small, rigid spines that give the female spikes a markedly bristly texture. In contrast, male inflorescences are fairly soft to the touch.

Blooming Palmer amaranth
Figure 2. Palmer amaranth in bloom, including male plants with anthers shedding pollen (center) and a female plant (upper right). Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Biology

In its native desert habitat, Palmer amaranth grows as a summer ephemeral herb supremely adapted to the rigors of intense heat and low, unpredictable rainfall (Ehleringer, 1983). Adaptive traits include the C4 photosynthetic pathway, a phenomenally high photosynthetic rate (even higher than most other C4 plants), optimum photosynthesis at leaf temperatures of 95–115 °F, capacity to continue photosynthesis under all but the most extreme drought stress, very high water use efficiency, and diurnal leaf movements that keep leaf blades perpendicular to the sun for maximum carbon fixation (Ibid.) . Additional traits include rapid seed germination, early seedling growth, and larger root volume than other amaranths (Steckel et al., 2004; Guo and Al-Khatib, 2003). Together, these traits allow Palmer amaranth to emerge, grow, and complete its life cycle on the soil moisture available at the time of germination (Ehleringer, 1983). In southern Arizona natural stands can attain dry weights of 2.2 tons per acre within 4 weeks after emergence (Ehleringer, 1983), which approaches the biomass of a mature winter annual cover crop.

In germination tests, Palmer amaranth seeds germinated rapidly—ithin 1–2 days—at a wide range of constant or alternating temperatures from 59–105 °F, with highest germination percentages and most rapid germination at 86–95 °F (Steckel et al, 2004; Guo and Al-Khatib, 2003). The primary requirement for germination seems to be moisture, as might be expected for a desert ephemeral. In the field, Palmer amaranth emergence occurs over an extended period (Jha et al., 2008b).

The seeds of Palmer amaranth have been reported to lose viability within 3 years when buried in the soil in Alabama and Georgia (Langcuster, 2008); however seed longevity in soil for the closely related redroot pigweed (A. retroflexus) and waterhemp (A. rudis) has been reported as short as 3–4 years in Mississippi and Illinois (Egley and Williams, 1990; Steckel et al., 2007) and as long as 12 years in Nebraska (Burnside et al., 1996). Thus, it is possible that some Palmer amaranth seeds remain viable in the soil for much longer than 3 years under certain conditions.

In field studies conducted in California (Keeley et al., 1987), Texas (Menges, 1988), Missouri (Sellers et al., 2003), Kansas (Horak, 1997; Horak and Loughin, 2000), and Arkansas (Fugate, 2009), Palmer amaranth has demonstrated a potential for extremely rapid growth and prolific seed set in cropland. It attained heights of 4 inches within 2–3 weeks after planting (WAP), and 35–40 inches at 5–7 WAP. For comparison, redroot pigweed and common waterhemp reached 2–3 inches at 2–3 WAP, and 20–30 inches at 5–7 WAP. Mature Palmer amaranth plants can reach heights of 6–10 ft with stems 2–3 inches thick (Fig. 3), forming 200–900 thousand mature seeds per female plant. Dry weight biomass of solid stands has been estimated as high as 5–9 tons per acre. Palmer amaranth considerably exceeded common waterhemp (Amaranthus rudis), redroot pigweed (A. retroflexus), and other Amaranthus species in height, dry weight, and leaf area in comparative growth analyses conducted under field conditions in Kansas (Horak and Loughin, 2000) and Missouri (Sellers et al, 2003). In growth chamber studies, Palmer amaranth grew more rapidly and formed larger root systems than redroot pigweed and common waterhemp in hot conditions (95 °F day, 86 °F night), and demonstrated the greatest heat tolerance and the least tolerance to cool conditions (Guo and Al-Khatib, 2003).

Palmer amaranth grows large
Figure 3. (a) Large specimen of Palmer amaranth, about 10 feet tall. (b) Stem of a mature Palmer amaranth. Photo credits: Rebekah D. Wallace, Bugwood.org.

Impact on Crops

The combination of rapid growth rate, adaptation to heat and drought, and large root volume makes Palmer amaranth an aggressive competitor against warm season crops (Fig. 4a), and a serious nuisance at harvest time (Fig. 4b). Once established, it can be very hard to control. In Georgia, some cotton farmers have resorted to manual pulling, as the weed has developed herbicide resistance, and regrows readily after chopping (Langcuster, 2008).

Palmer amaranth is aggressive
Figure 4. (a) A vigorous, much-branched Palmer amaranth has displaced the soybean crop from several feet of row. (b) Palmer amaranth in cotton at crop maturity interferes with harvest. Photo credit: (a) Rebekah D. Wallace, Bugwood.org; (b) Joseph LaForest, University of Georgia, Bugwood.org.

Palmer amaranth causes significant yield reductions in all agronomic row crops, especially when it emerges before or with the crop. In a field study in Arkansas, one Palmer amaranth per 10 ft of row reduced soybean grain yield by 17%, and one weed per foot of row cut yields 64% when crop and weeds emerged together (Klingman and Oliver, 1994). The amaranth exceeded the crop in height by 8–24 inches from 4 weeks after emergence through harvest. Corn yields were reduced 20% by one Palmer amaranth per 6.6 feet of row, and 40–80% by one weed per foot of row in Kansas (Massinga et al., 2001). Amaranth height exceeded that of corn, and its foliage intercepted light at a greater height above the ground than corn foliage (Massinga et al, 2003). However, when the weed emerged several weeks after corn, it had much less impact on yield, and its seed production was reduced by 80–98% (Massinga et al., 2001). In another Kansas field trial, Palmer amaranth planted with soybean reduced crop yield 28%, whereas Palmer amaranth planted 15–20 days after soybean had no effect on crop yield (Bensch et al., 1997).

Residues of Palmer amaranth can suppress crop growth. Greenhouse and field studies indicate that incorporation of a heavy stand of Palmer amaranth into the soil just before planting can significantly hinder seedling growth in carrot, onion, cabbage, and grain sorghum (Menges 1987, 1988), and the authors suggested that allelopathy (release of natural plant growth inhibitors from the residues) may play a role in this effect. The growth of Palmer amaranth itself may be retarded somewhat by allelochemicals from cover crops in the Brassica (mustard) family. Isothiocyanate compounds derived from Brassica residues reduced Palmer amaranth emergence in greenhouse trials (Norsworthy and Meehan, 2005). Incorporation of the cover crops themselves into field soil prior to planting pepper reduced Palmer amaranth levels by 25–50% during the first four weeks in one year out of two (Norsworthy et al., 2007).

Management Implications for Organic Production

Palmer amaranth is clearly the most aggressive pigweed in hot, humid to semiarid conditions. Organic producers in the southern half of the U.S. are well advised to get a positive identification on pigweeds to determine whether this species is present.

Like other pigweeds, Palmer amaranth is quite vulnerable to cultivation during the seedling stage, but its unusually rapid early development leaves a shorter time window for control. Diligent monitoring and timely intervention are critical for the control of Palmer amaranth, as cultivation and flaming are most effective on weeds not more than 1 inch tall.

The temperature optimum for Palmer amaranth growth is higher than that of most vegetable and row crops. Similarly, its drought tolerance is greater than that of most cultivated crops. In cooler conditions with adequate moisture, the weed may lose its competitive edge against most crops. Therefore, planting dates may be a significant factor in managing Palmer amaranth; for example, frost-tender vegetables like tomato or snap bean may be grown in spring or fall in the Gulf Coast states, when moderate temperatures favor the vegetable over the weed.

Although Palmer amaranth seeds may have limited longevity in the soil in hot, rainy climates (Langcuster, 2008), it is especially important to prevent seed production by this weed in order to draw down the seed bank. In intensive vegetable production, it is worth the effort to pull out any Palmer amaranth individuals that escape cultivation before they set seed. If a heavy seed shed of Palmer amaranth occurs, inversion tillage may be useful in limiting weed emergence in the following season; however, additional inversion should be avoided for the next several years so that viable Palmer amaranth seeds are not brought back to the surface.

A diversified crop rotation that varies tillage, planting, and harvest schedules from year to year as well as crop species and plant family, can help reduce problems with summer annual weeds, and may be helpful in managing Palmer amaranth. Incorporating a radish, mustard, or other brassica green manure may help slow emergence and growth of Palmer amaranth; however brassica allelopathy should not be counted on to control the weed. Caution must also be taken to avoid suppressing crop germination, emergence, and growth by brassica residues, especially in direct-sown small-seeded vegetables and peas.

The organic weed management techniques outlined in the general article on pigweeds are appropriate. Additional recommendations for fields with significant populations of Palmer amaranth include:

  • After planting, scout every 2–3 days for weed emergence.
  • When pigweed seedlings are detected, cultivate or flame immediately – don't wait until you can determine whether they are Palmer amaranth.
  • If practical, adjust planting dates to avoid weed–crop competition during very hot weather.
  • To reduce heavy infestations, rotate to cool season production crops, and focus on weed control through timely tillage and cover cropping during summer months.
References Cited
  • Bensch, C. J., M. J. Horak, and D. E. Peterson. 1997. Competition of three Amaranthus species in soybean. North Central Weed Science Society Proceedings 52: 148.
  • Burnside, O. C., R. G. Wilson, S. Weisberg, and K. G. Hubbard. 1996. Seed longevity of 41 weed species buried 17 years in eastern and western Nebraska. Weed Science 44: 74–86. (Available online at: http://www.jstor.org/stable/4045786) (verified 10 Sept 2012).
  • Culpeper, A. S., T. L. Grey, W. K. Vencill, J. M. Kitchler, T. M. Webster, S. M. Brown, A. C. York, J. W. Davis, and W. W. Hanna. 2006. Glyphosate-resistant Palmer amaranth (Amaranthus palmeri) confirmed in Georgia. Weed Science 54: 620–626. (Available online at: http://dx.doi.org/10.1614/WS-06-001R.1) (verified 10 Sept 2012).
  • Egley, G. H., and R. D. Williams. 1990. Decline of weed seeds and seedling emergence over five years as affected by soil disturbances. Weed Science 38: 504–510. (Available online at: http://www.jstor.org/stable/4045064) (verified 10 Sept 2012).
  • Ehleringer, J. 1983. Ecophysiology of Amaranthus palmeri, a Sonoran desert summer annual. Oecologia 57: 107–112. (Available online at: http://dx.doi.org/10.1007/BF00379568) (verified 10 Sept 2012).
  • Fugate, L. 2009. Pigweed causing farmers to rethink farming methods. University of Arkansas Division of Agriculture Cooperative Extension Service News - October 2009.
  • Guo, P., and K. Al-Khatib. 2003. Temperature effects on germination and growth of redroot pigweed (Amaranthus retroflexus), Palmer amaranth (A. palmeri), and common waterhemp (A. rudis). Weed Science 51: 869–875. (Available online at: http://dx.doi.org/10.1614/P2002-127) (verified 10 Sept 2012).
  • Horak, M. J. 1997. The changing nature of palmer amaranth: A case study. North Central Weed Science Society Proceedings 52: 161.
  • Horak, M. J., and T. M. Loughin. 2000. Growth analysis of four Amaranthus species. Weed Science 48: 347–355. (Available online at: http://dx.doi.org/10.1614/0043-1745(2000)048%5B0347:GAOFAS%5D2.0.CO;2) (verified 10 Sept 2012).
  • Horak, M. J., and D. E. Peterson. 1995. Biotypes of Palmer amaranth (Amaranthus palmeri) and common waterhemp (Amaranthus rudis) are resistant to imazethapyr and thifensulfuron. Weed Technology 9: 192–195. (Available online at: http://www.jstor.org/stable/3987844) (verified 10 Sept 2012).
  • Jha, P., J. K. Norsworthy, W. Bridges, Jr., and M. B. Riley. 2008a. Influence of glyphosate timing and row width on Palmer amaranth (Amaranthus palmeri) and pusley (Richardia spp.) demographies in glyphosate-resistant soybean. Weed Science 56: 408–415. (Available online at: http://dx.doi.org/10.1614/WS-07-174.1) (verified 10 Sept 2012).
  • Jha, P., J. K. Norsworthy, M. B. Riley, D. G. Bielenberg, and W. Bridges, Jr. 2008b. Acclimation of Palmer amaranth (Amaranthus palmeri) to shading. Weed Science 56: 729–734. (Available online at: http://dx.doi.org/10.1614/WS-07-203.1) (verified 10 Sept 2012).
  • Keeley, P. E., C. H. Carter, and R. J. Thullen. 1987. Influence of planting date on growth of Palmer amaranth (Amaranthus palmeri). Weed Science 35: 199–204. (Available online at: http://www.jstor.org/stable/4044391) (verified 10 Sept 2012).
  • Klingman, T. E., and L. R. Oliver. 1994. Palmer amaranth (Amaranthus palmeri) interference in soybeans (Glycine max). Weed Science 42: 523–527. (Available online at: http://www.jstor.org/stable/4045448) (verified 10 Sept 2012).
  • Langcuster, J. 2008. Scarier than Halloween – the nightmare weed that threatens Southern row crops. Extension Daily, Alabama Cooperative Extension, October 22, 2008. (Available online at: http://www.aces.edu/department/extcomm/npa/daily/archives/003801.php) (verified 10 Sept 2012).
  • Massinga, R. A., R. S. Currie, M. J. Horak, and J. Boyer, Jr. 2001. Interference of Palmer amaranth in corn. Weed Science 49: 202–208. (Available online at: http://dx.doi.org/10.1614/0043-1745(2001)049%5B0202:IOPAIC%5D2.0.CO;2) (verified 10 Sept 2012).
  • Massinga, R. A., R. S. Currie, and T. P. Trooien. 2003. Water use and light interception under Palmer amaranth (Amaranthus palmeri) and corn competition. Weed Science 51: 523–531. (Available online at: http://dx.doi.org/10.1614/0043-1745(2003)051%5B0523:WUALIU%5D2.0.CO;2) (verified 10 Sept 2012).
  • Menges, R. M. 1987. Allelopathic effects of Palmer amaranth (Amaranthus palmeri) and other plant residues in soil. Weed Science 35: 339–347. (Available online at: http://www.jstor.org/stable/4044595) (verified 10 Sept 2012).
  • Menges, R. M. 1988. Allelopathic effects of Palmer amaranth (Amaranthus palmeri) on seedling growth. Weed Science 36: 325–328. (Available online at: http://www.jstor.org/stable/4044643) (verified 10 Sept 2012).
  • Norsworthy, J. K., M. S. Malik, P. Jha, and M. B. Riley. 2007. Suppression of Digitaria sanguinalis and Amaranthus palmeri using autumn-sown glucosinolate-producing cover crops in organically grown pepper. Weed Research 47: 425–432. (Available online at: http://dx.doi.org/10.1111/j.1365-3180.2007.00586.x) (verified 10 Sept 2012).
  • Norsworthy, J. K., and J. T. Meehan, IV. 2005. Use of isothiocyanates for suppression of Palmer amaranth (Amaranthus palmeri), pitted morningglory (Ipomoea lacunose), and yellow nutsedge (Cyperus esculentus). Weed Science 53: 884–890. (Available online at: http://dx.doi.org/10.1614/WS-05-056R.1) (verified 10 Sept 2012).
  • Sellers, B. A., R. J. Smeda, W. G. Johnson, J. A. Kendig, and M. R. Ellersieck. 2003. Comparative growth of six Amaranthus species in Missouri. Weed Science 51: 329–333. (Available online at: http://dx.doi.org/10.1614/0043-1745(2003)051%5B0329:CGOSAS%5D2.0.CO;2) (verified 10 Sept 2012).
  • Steckel, L. E., C. L. Sprague, E. W. Stoller, and L. M. Wax. 2004. Temperature effects on germination of nine Amaranthus species. Weed Science 52: 217–221. (Available online at: http://dx.doi.org/10.1614/WS-03-012R) (verified 10 Sept 2012).
  • Steckel, L. E., C. L. Sprague, E. W. Stoller, L. M. Wax, and F. W. Simmons. 2007. Tillage, cropping system, and soil depth effects on common waterhemp (Amaranthus rudis) seed-bank persistence. Weed Science 55: 235–239. (Available online at: http://dx.doi.org/10.1614/WS-06-198) (verified 10 Sept 2012).
  • Webster, T. M. 2006. Weed survey – southern states. Vegetable, fruit and nut crops subsection. Proceedings of the Southern Weed Science Society 59: 260–277. (Available online at: http://www.swss.ws/NewWebDesign/Publications/Weed%20Survey%20Archives/Southern%20Weed%20Survey%202006%20Vegetables%20and%20Fruits.pdf) (verified 10 Sept 2012).

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 5122

Case Studies on Organic Weed Management

mer, 2012/09/12 - 16:53

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T1046

Purple Nutsedge (Cyperus rotundus) in Greater Depth

mer, 2012/09/12 - 16:26

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Introduction

Purple nutsedge (Cyperus rotundus) is a colony-forming perennial weed that seriously impacts agriculture across the southernmost United States. Native to tropical Eurasia, purple nutsedge has become a major weed of vegetable, row, and plantation crops in tropical and warm temperate climates around the world, is very difficult to manage with either organic or conventional weed control strategies (William, 1976; Bangarwa et al., 2008; Wang et al., 2008), and has been called the world's worst weed (Holm et al., 1991). Purple nutsedge is one of the most extensively researched non-cultivated plant species on the planet, yet the complexities of its life cycle, and its multiple adaptations to environmental extremes and weed control tactics are as yet incompletely understood.

Purple nutsedge is a grass-like weed in the sedge family (Cyperaceae) with top growth 4–30 inches tall (Fig. 1), an extensive underground network of basal bulbs, fibrous roots, thin wiry rhizomes (Fig. 2), and tubers borne in chains of 2–6 or more on rhizomes, with tubers spaced 2-10 inches apart. The leaves are mostly basal, dark green, 0.1–0.25 inches wide with a prominent midrib, and abruptly tapered at the tips. The purplish to red-brown inflorescence (Fig. 1) is borne on a culm (stem) that is triangular in cross section and usually taller than the foliage (Bryson and DeFelice, 2009). The inflorescence itself consists of an umbel of spikes, some of which are sessile, and others are borne on stalks of unequal length.The subtending leaflike bracts are usually shorter than the longest spikes.

Purple nutsedge in bloom
Figure 1. Purple nutsedge in bloom, showing purplish color of flower heads. Photo credit: Forest & Kim Starr, U.S. Geological Survey, Bugwood.org.

Purple nutsedge clone
Figure 2. Several purple nutsedge plants linked by a network of rhizomes. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Biology

Purple nutsedge initiates its seasonal growth cycle almost entirely from tubers, as viable seeds rarely occur in this species. Tuber dormancy is broken by high temperature (90-100 °F) and diurnal temperature fluctuations. In Hawaii populations, 30 minute daily pulses of 95 °F over a baseline of 68 °F stimulated shoot emergence from tubers as effectively as continuous 95 °F, and an alternating regime of 65 °F and 75 °F resulted in more emergence than constant 75 °F (Miles et al., 1996). This response to temperature fluctuation promotes nutsedge emergence in the absence of a shading canopy. Chilling has been reported to promote tuber sprouting (Shamsi et al., 1978), but tubers are killed by freezing temperatures (Holm et al., 1991).

Each tuber has multiple buds, most of which remain dormant and act as a reserve in the event that the active shoot is destroyed. Tuber chains show apical dominance, so that the terminal tuber initiates active growth while many or all of the others on the chain remain dormant unless the terminal tuber is destroyed or the chain is broken (Kawabata and Nishimoto, 2003). Dormant tubers commonly persist in the soil for 3–4 years, and can remain viable for as long as 10 years in some conditions (California Department of Food and Agriculture)

The tuber sprout consists of a sharp pointed rhizome, which grows toward the soil surface, then differentiates into shoot and leaves in response to light (Chase et al., 1998). The plant forms a subterranean basal bulb, which contains the shoot meristem (site of cell division and formation of new leaves). Basal bulbs form mostly within 3 inches of the soil surface, although bulbs have been observed at 4–8 inches (Hauser, 1962; Holm et al., 1991; William, 1976; William and Warren, 1975) (Fig. 3). Bulbs develop fibrous root systems that may extend 4 feet deep in the soil profile (Holm et al., 1991; California Department of Food and Agriculture). Because the shoot growing point remains in the basal bulb, leaves regrow readily if severed at the soil surface (William and Warren, 1975).

Purple nutsedge depth of bulbs
Figure 3. Most of the basal bulbs in this purple nutsedge stand are within 1.5 inches of the surface, but one bulb can be seen sending up a shoot from a depth of about 4 inches. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Within 2–3 weeks after shoot emergence, basal bulbs send out new rhizomes that form additional bulbs and daughter plants (Fig. 4). The cycle repeats several times during a growing season, so that a single sprouting tuber can give rise to hundreds of shoots. In a field trial in the low desert of southeastern California, purple nutsedge planted at a density of about 0.07 tuber per square foot and left uncontrolled increased to about 22 tubers per square foot at the end of one season, and 115 tubers per square foot at the end of the second season (Wang et al., 2008).

Purple nutsedge with mother tubers
Figure 4. Young purple nutsedge plants, still attached to their mother tubers by rhizomes ranging from a fraction of an inch (left) to several inches in length (right), depending on the depth of the tuber in the soil. Two of the plants have initiated new rhizomes and daughter plants. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Plants usually flower about 7–8 weeks after emergence, although flowering can occur as early as 3 weeks. New tubers begin to form at about the time of flowering. Most tubers are set in the top 8 inches of the soil profile, with a few forming at greater depths. After flowering, purple nutsedge undergoes a marked shift from aboveground to belowground development, so that tubers continue to form for several weeks after shoot growth ceases. In temperate latitudes, this cycle corresponds with the seasons, and tuber formation begins in late summer as photoperiods become shorter (Jordan-Molero and Stoller, 1978). However purple nutsedge populations in Costa Rica and Brazil showed similar developmental cycles (Neeser et al., 1997; William, 1976); a new cycle may be triggered by soil preparation at the beginning of a cropping season (William and Warren, 1975).

In a field trial in Georgia, purple nutsedge grown in fertilized soil without competing vegetation flowered and began to form tubers about 7–8 weeks after emergence, and developed 2.4 tons above ground and 4.2 tons below ground dry weight per acre by 12 weeks (Hauser, 1962). After that point, active foliar growth diminished while tuber formation accelerated; by 20 weeks, shoot biomass was 3.3 tons per acre while below ground biomass reached 12 tons per acre. Underground dry weights of approximately 5–8 tons per acre have been observed in naturally occurring heavy infestations at other sites (Holm et al., 1991). For comparison, the above ground biomass of an 8-ft pearl millet or sorghum-sudangrass cover crop with a fully closed canopy is about 5–6 tons per acre.

Purple nutsedge has the C4 photosynthetic pathway, which contributes to its ability to grow and spread rapidly in hot weather and high light levels. The weed shows tremendous heat tolerance in field conditions, yet tubers can be killed either by desiccation to 15–24% moisture content in direct sun, or by exposure to 122 °F for 12 hours (Holm et al., 1991; Webster, 2003). However, tubers and bulbs located several inches deep in the soil profile are shielded from lethal temperatures, and the deep, fibrous root system keeps tubers hydrated. When drought, flooding, or other unfavorable conditions occur, foliage dies back and viable dormant tubers remain.

Like most C4 plants, purple nutsedge is shade intolerant, and can be suppressed by a closed crop canopy, although tubers remain viable and send up new shoots when the canopy is removed (Holm et al., 1991). In greenhouse trials, neutral shade (white cheesecloth) that reduced incident light by 20% reduced purple nutsedge growth (dry weight accumulation) by 25%, whereas 60% shade cut aboveground dry weight by 80% and tuber dry weight by 97% (Santos et al., 1997b). In a field trial in Brazil, nutsedge compensated for light reductions by shade cloth up to 63% by increasing leaf length and plant height, with no decrease in biomass (William and Warren, 1975). However, a similar degree of shade from crop canopies greatly reduced new tuber production by purple nutsedge during the cropping cycle in Costa Rica (Neeser et al., 1997). Bush snap bean, a bush bean/sweet corn intercrop, and sweet potato competed more effectively than corn alone, pole bean, or bell pepper.

In India, native soil endomycorrhizal fungi were found to colonize purple nutsedge, but failed to form the mutually beneficial arbuscular structures in the plant's roots, and significantly reduced nutsedge growth rates (Muthukumar et al., 1997). When onion, a “nurse plant” that forms a beneficial symbiosis with the same fungi, was grown with nutsedge, the adverse impacts of mycorrhizae on nutsedge growth were accentuated. Keeping the soil flooded inhibited mycorrhizal development and restored nutsedge growth to that of mycorrhizae-free controls.

A few crops may retard purple nutsedge growth through allelopathy. Four foliar applications of a water extract of sorghum (containing water soluble allelochemicals) significantly reduced growth of a weed flora dominated by purple nutsedge, and protected corn yields more economically than either hand weeding or the herbicide pendimethalin (Cheema et al., 2004). In greenhouse studies, sweet potato cv. ‘Regal’ reduced purple nutsedge growth through allelopathy, although the crop was similarly suppressed by the nutsedge (Peterson and Harrison, 1995). In field trials, Neeser et al. (1997) found that sweet potato suppressed purple nutsedge tuber formation to a substantially greater degree than can be attributed to shading alone, and hypothesized that allelopathic effects contributed to the suppression. A 4-ton-per-acre cowpea cover crop, mowed at maturity and left as a mulch, suppressed weed growth (of which purple nutsedge was a major component) by 70–90% during pepper production in the desert of southern California (Hutchinson and McGiffen, 2000). Since nutsedge readily penetrates organic mulch, an allelopathic effect of cowpea on the weed may be involved. On the other hand, incorporation of a turnip cover crop (rich in isothiocyanates, a potent class of allelochemicals in brassicas) failed to affect purple nutsedge growth in South Carolina field trials (Bangarwa et al., 2008).

Impacts on Crop Production

Purple nutsedge competes vigorously against most crops for soil moisture and nutrients (Fig. 5), and against low-growing or slow-starting crops for light. It is especially competitive during warm seasons with ample moisture, and becomes less so in cooler, drier conditions (William and Warren, 1975). Purple nutsedge sometimes occurs with the closely related weed yellow nutsedge; however purple nutsedge is the more aggressive of the two in hot climates (Wang et al., 2008), whereas the reverse is true in cooler conditions (Jordan-Molero and Stoller, 1978).

High levels of available nutrients and moisture seem to intensify purple nutsedge competition against crops. Vegetable farmers in Brazil and Panama reduce nutsedge competition by placing fertilizer and water near crop plants rather than applying water and nutrients to the entire field (William, 1976). Increasing fertilizer N rates have accentuated yield losses to purple nutsedge in field trials with radish (Santos et al., 1998), tomato (Morales-Payan et al., 1997) and upland rice (Okafor and De Datta, 1976). Greenhouse trials have suggested a similar pattern in the interaction between purple nutsedge and pepper (Morales-Payan et al., 1998) and cilantro (Morales-Payan et al., 1999), and increased nutsedge competitiveness toward cotton at higher soil moisture levels (Cinco-Castro and McCloskey, 1997).

Purple nutsedge in peppers
Figure 5. A heavy infestation of nutsedge within crop rows has seriously affected this pepper crop by competing for moisture. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Substances released from living or decaying below-ground parts of purple nutsedge have shown allelopathic activity against barley, mustard, and cotton (Friedman and Horowitz, 1971; Horowitz and Friedman, 1971), and sweet potato (Peterson and Harrison, 1995) in greenhouse trials. Given the large underground biomass in a heavy purple nutsedge infestation, the possible contribution of allelopathy to this weed's adverse impacts on crops many times its height in field conditions merits further investigation.

Purple nutsedge readily penetrates black plastic film mulch (Webster, 2005a) (Fig. 6). Black plastic doubled the rate at which purple nutsedge spread and propagated in field studies in Georgia (Webster, 2005b), probably because the mulch maintained higher soil temperatures. However, clear or translucent plastic film mulches reduce purple nutsedge growth because the emerging shoots open their leaves in response to light before penetrating the film, and become trapped (Patterson, 1998; Chase et al., 1998).

Purple nutsedge emerging through black plastic
Figure 6. Purple nutsedge emerging through black plastic film mulch. Photo credit: Rebekah D. Wallace, Bugwood.org.

Management Strategies and Tactics for Organic Production

Mechanical control of an invasive perennial weed infestation begins with an initial vigorous tillage to fragment the weed, followed by additional cultivations whenever fragments have regenerated new shoots with 3–4 leaves, at which time the weed's underground reserves have been drawn down to their lowest point (Mohler and DiTommaso, unpublished). For mechanical control of nutsedge, repeating cultivation every 2–3 weeks, before plants reach the 6-leaf stage (Fig. 7) has been recommended (California Department of Food and Agriculture; Russ and Burgess, 2009).

Young purple nutsedge
Figure 7. The purple nutsedge regrowth in this photo has about 56 leaves per plant, and will begin rebuilding reserves and forming new daughter plants unless it is cultivated immediately. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Purple nutsedge is difficult to control through this strategy because of its tremendous underground reserves and because shoot growing points remain below ground in the basal bulb. In South Carolina, the weed has been observed to regenerate substantial shoot growth and begin forming new rhizomes within 2 weeks after tillage (Fig. 8).Experiments with clipping top growth every 2 weeks for 8 months, or every 6 days for 6 weeks weakened but did not kill tubers (Santos et al., 1997a; William, 1976). In field trials cultivation every 2–3 weeks for 2 years has reduced tuber populations by 80% (Holm et al., 1991). However, such intensive cultivation degrades soil quality, and may not be practical. Thus, cultivation must be used in conjunction with other tactics for effective nutsedge control.


Purple nutsedge regrowth 13 days after tillage
Figure 8. (a) Purple nutsedge regrowth just 13 days after field in the South Carolina piedmont was rototilled to a depth of 4 inches, bedded, and planted to sweet potato. Alleys were tilled again a few days prior to the photograph. (b) Individual plants forming new rhizomes 13 days after tillage. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

In the humid parts of the southern U.S., soil solarization—heating the soil by covering with clear or translucent film during hot sunny weather—does not produce lethal temperatures deep enough in the soil profile to eradicate nutsedge tubers (Webster, 2003), and may even stimulate tuber sprouting and shoot emergence (Egley, 1983). However, sprouting tubers are weakened when the emerging foliage is trapped and heat-killed under the film. Substantial reduction of purple nutsedge infestations has been achieved in field trials in northern Florida, especially under a thermal-infrared-retentive (TIR) plastic film that gives more intense heating than clear polyethylene (Chase et al., 1999).

In the Coachella Valley (low desert region) of southeastern California, where average daily maximum temperatures in July exceed 104 °F, solarization with black plastic during a summer fallow period generated temperatures lethal to nutsedge tubers to a depth of 6 inches (Wang et al., 2008) This treatement virtually eliminated purple nutsedge in a subsequent fall broccoli crop. In the same trial, repeated manual cultivation reduced tuber populations by 93%, yet nutsedge competition still slashed broccoli yield 80%. A summer smother crop of sudangrass proved ineffective in reducing nutsedge populations.

Poultry and hogs consume nutsedge tubers, and have shown potential for purple nutsedge control. Early experiments in Alabama with running laying hens in fields with purple nutsedge infestations of about 20 tubers per square foot showed that a stocking rate of 480 birds per acre maintained for one full growing season eradicated the weed (Mayton et al., 1945). However, since chickens forage mostly within 50 feet of the henhouse, enclosing 300 chickens in a single 0.5-acre pen was not effective in cleaning the whole area. Chickens used for this purpose should be one of the more actively-foraging breeds, and must be acclimiatized to grazing. Mayton et al. (1945) also found that weeder geese at 4 to 16 birds per 0.5-acre could clean purple nutsedge out of a cotton field, but geese were less effective in cleaning up a fallow field. Tillage to break up tuber chains (thus breaking dormancy) enhanced efficacy of nutsedge eradication by geese.

In India, pigs are sometimes used to remove purple nutsedge from rice paddies before planting the crop. Pigs readily root up and consume the tubers, and running 60-75 animals in a 2.5 acre (1 hectare) field for one day has been reported to provide effective control (OSWALD, 1997).

On USDA-certified organic farms, livestock and poultry must be removed from the field and their droppings incorporated into the soil at least 120 days before harvest of an organic food crop that may come into contact with soil or soil particles, or 90 days for a crop not so exposed, such as tree fruit or sweet corn.

Efforts to develop integrated organic management strategies for purple nutsedge have thus far been only partially successful. In Clemson, South Carolina, March–July fallow treatments of solarization with clear or green translucent film, a turnip cover crop followed by solarization, or tillage every 3 weeks were followed by hand weeding, straw mulching, or no weed control during fall pepper production. Two successive years of fallow treatments followed by hand weeding reduced purple nutsedge tuber populations by 36–58%, whereas straw mulch was less effective, and tuber populations increased substantially without weed control during pepper production (Bangarwa et al., 2008). The authors concluded: “when selecting a site for organic crop production, an effort should be made to select one free of purple nutsedge”.

In Gainesville, Florida, researchers and farmers are testing a rotation of fall vegetables (lettuce or broccoli), spring vegetables (pepper or squash), and summer fallow with repeated tillage, repeated flaming, cover crop, or solarization. Treatments reduced purple nutsedge populations by approximately half during the first year, but little further reduction occurred during the second year, when high summer rainfall favored the weed (Koenig and Chase, 2010).

Highly competitive cover or cash crops such as jack bean, velvet bean, soybean, cotton, and chayote, have been used with some success against purple nutsedge in Brazil and other tropical regions (William, 1976). Competitive vegetable crops, including bush bean, cucumber, and transplanted cabbage may require only one hand weeding 3–5 weeks after planting (critical weed-free period) to prevent yield loss to purple nutsedge (William and Warren, 1975). Bush snap bean and sweet potato grown without any post-emergence weed control almost stopped tuber propagation during the cropping cycle in Costa Rica (Neeser et al., 1997).

Efforts continue to develop effective organic management strategies for purple nutsedge. Successful strategies will likely integrate several key components:

  • Break tuber dormancy by physically breaking up tuber chains and providing thermal stimuli for germination.
  • Disrupt emerging new growth, exhaust tuber reserves, and prevent propagation through timely cultivation. If practical, cultivate deep enough to sever basal bulbs from root systems and mother tubers.
  • Provide strong crop competition, with crop canopies generating at least 80% shade during critical times for nutsedge suppression, such as during tuber set in late summer.
  • Manage nutrients and moisture to favor the crop over the nutsedge; avoid overwatering and overfertilizing.

Additional tactics may include soil solarization in very hot sunny climates; weed removal by hogs,poultry, or geese; shifting crop planting and harvest dates to avoid the most intense nutsedge competition; encouraging soil mycorrhizal fungi; and growing crops known to be allelopathic to nutsedge.

References Cited
  • Bangarwa, S. K., J. K. Norsworthy, P. Jha, and M. Malik. 2008. Purple nutsedge (Cyperus rotundus) management in an organic production system. Weed Science 56: 606–613. (Available online at: http://dx.doi.org/10.1614/WS-07-187.1) (verified 10 Sept 2012).
  • Bryson, C. T., and M. S. DeFelice. 2009. Weeds of the South. University of Georgia Press, Athens, GA.
  • California Department of Food and Agriculture. Weed Information – Yellow Nutsedge and Purple Nutsedge. (Available online at: http://www.cdfa.ca.gov/plant/ipc/weedinfo/cyperus.htm) (verified 10 Sept 2012).
  • Chase, C. A., T. R. Sinclair, D. G. Shilling, J. P. Gilreath, and S. J. Locascio. 1998. Light effects on rhizome morphogenesis in nutsedges (Cyperus spp): Implications for control by soil solarization. Weed Science 46: 575–580. (Available online at: http://www.jstor.org/stable/4045964) (verified 10 Sept 2012).
  • Chase, C. A., T. R. Sinclair, and S. J. Locascio. 1999. Effects of soil temperature and tuber depth on Cyperus spp. control. Weed Science 47: 467–472. (Available online at: http://www.jstor.org/stable/4046223) (verified 10 Sept 2012).
  • Cheema, Z. A., A. Khaliq, and S. Saeed. 2004. Weed control in maize (Zea mays L.) through sorghum allelopathy. Journal of Sustainable Agriculture 23: 73–87. (Available online at: http://dx.doi.org/10.1300/J064v23n04_07) (verified 10 Sept 2012).
  • Cinco-Castro, R., and W. B. McCloskey. 1997. Purple nutsedge (Cyperus rotundus L) competition with cotton under wet and dry soil moistures. Weed Science Society of America Abstracts 37: 55.
  • Egley, G. H. 1983. Weed seed and seedling reductions by soil solarization with transparent polyethylene sheets. Weed Science 31: 404–409. ((Available online at: http://www.jstor.org/stable/4043730) (verified 10 Sept 2012).
  • Friedman, T., and M. Horowitz. 1971. Biologically active substances in subterranean parts of purple nutsedge. Weed Science 19: 398–401. (Available online at: http://www.jstor.org/stable/4041790) (verified 10 Sept 2012).
  • Hauser, E. W. 1962. Development of purple nutsedge under field conditions. Weeds 10: 315–321. (Available online at: http://www.jstor.org/stable/4040836) (verified 10 Sept 2012).
  • Holm, L. G., D. L. Plucknett, J. V. Pancho, and J. P. Herberger, 1991. The world's worst weeds. Kriegar Publishing Company, Malabar, FL.
  • Horowitz, M., and T. Friedman. 1971. Biological activity of subterranean residues of Cynodon dactylon L., Sorghum halapense L., and Cyperus rotundus L. Weed Research 11: 88–93. (Available online at: http://dx.doi.org/10.1111/j.1365-3180.1971.tb00982.x) (verified 10 Sept 2012).
  • Hutchinson, C. M., and M. E. McGiffen, Jr. 2000. Cowpea cover crop mulch for weed control in desert pepper production. HortScience 35: 196–198. (Available online at: http://hortsci.ashspublications.org/content/35/2/196.abstract) (verified 10 Sept 2012).
  • Jordan-Molero, J. E., and E. W. Stoller. 1978. Seasonal development of yellow and purple nutsedges (Cyperus esculentus and C. rotundus) in Illinois. Weed Science 26: 614–618. (Available online at: http://www.jstor.org/stable/4042940) (verified 10 Sept 2012).
  • Kawabata, O., and R. K. Nishimoto. 2003. Temperature and rhizome chain effect on sprouting of purple nutsedge (Cyperus rotundus) ecotypes. Weed Science 51: 348–355. (Available online at: http://dx.doi.org/10.1614/0043-1745(2003)051[0348:TARCEO]2.0.CO;2) (verified 10 Sept 2012).
  • Koening, R., and C. A. Chase. 2010. Weed management techniques that really work. Presentation at the Southern Sustainable Agriculture Working Group Conference in Chattanooga, TN, January 22–23, 2010. Findings to be published in refereed journal.
  • Mayton, E. L., E. V. Smith, and D. King. 1945. Nutgrass eradication studies IV: use of chickens and geese in the control of nutgrass, Cyperus rotundus L. Journal of the American Society of Agronomy 37: 785–791.
  • Miles, J. E., R. K. Nishimoto, and O. Kawabata. 1996. Diurnally alternating temperatures stimulate sprouting of purple nutsedge (Cyperus rotundus) tubers. Weed Science 44: 122–125. (Available online at: http://www.jstor.org/stable/4045792) (verified 10 Sept 2012).
  • Mohler, C. A., and A. DiTommaso. Unpublished. Manage weeds on your farm: A guide to ecological strategies. Department of Crop and Soil Sciences, Cornell University. Pre-publication draft, version 5.1 (2008). Publication through SARE expected in 2012.
  • Morales-Payan, J. P., B. M. Santos, W. M. Stall, and T. A. Bewick. 1998. Interference of purple nutsedge (Cyperus rotundus) population densities on bell pepper (Capsicum annuum) yield as influenced by nitrogen. Weed Technology 12: 230–234. (Available online at: http://www.jstor.org/stable/3988380) (verified 10 Sept 2012).
  • Morales-Payan, J. P., B. M. Santos, W. M. Stall, and T. A. Bewick. 1999. Influence of nitrogen fertilization on the competitive interactions of cilantro (Coriandrum sativum) and purple nutsedge (Cyperus rotundus L.). Journal of Herbs, Spices and Medicinal Plants 6: 59–66. (Available online at: http://gcrec.ifas.ufl.edu/Weed%20Science/Documents/Weed%20Ecology%20Studies/JHMP9901.pdf) (verified 10 Sept 2012).
  • Morales-Payan, W. M. Stall, D. G. Shilling, J. A. Dusky, and T. A. Bewick. 1997. Influence of nitrogen on the interference of purple and yellow nutsedge (Cyperus rotundus and Cyperus esculentus) with tomato (Lycopersicon esculentum). HortScience 32: 431. (Available online at: http://hortsci.ashspublications.org/content/32/3/431.3.abstract) (verified 10 Sept 2012).
  • Muthukumar, T., K. Udaiyan, A. Karthikeyan, and S. Manian. 1997. Influence of native endomycorrhiza, soil flooding and nurse plant on mycorrhizal status and growth of purple nutsedge (Cyperus rotundus L.). Agriculture, Ecosystems and Environment 61: 51–58. (Available online at: http://dx.doi.org/10.1016/S0167-8809(96)01073-0) (verified 10 Sept 2012).
  • Neeser, C., R. Aguero, and C. J. Swanton. 1997. Incident photosynthetically active radiation as a basis for integrated management of purple nutsedge (Cyperus rotundus). Weed Science 45: 777–783. (Available online at: http://www.jstor.org/stable/4045844) (verified 10 Sept 2012).
  • Okafor, L. I., and S. K. De Datta. 1976. Competition between upland rice and purple nutsedge for nitrogen, moisture, and light. Weed Science 24: 43–46. (Available online at: http://www.jstor.org/stable/4042494) (verified 10 Sept 2012).
  • Open Source for Weed Assessment in Lowland Paddy Fields (OSWALD). 1997. Cyperus rotundus - Cyperacea. (Available online at: http://www.oswaldasia.org/species/c/cypro/cypro_en.html) (verified 10 Sept 2012).
  • Patterson, D. T. 1998. Suppression of purple nutsedge (Cyperus rotundus) with polyethylene film mulch. Weed Technology 12: 275–280. (Available online at: http://www.jstor.org/stable/3988388) (verified 10 Sept 2012).
  • Peterson, J. K., and H. F. Harrison. 1995. Sweet potato allelopathic substance inhibits growth of purple nutsedge. Weed Technology 9: 277–280. (Available online at: http://www.jstor.org/stable/3987745) (verified 10 Sept 2012).
  • Russ, K., and C. Burgess. 2009. Nutsedge. Clemson University Extension. 5 pp. (Available online at: http://www.clemson.edu/extension/hgic/pests/weeds/hgic2312.html) (verified 10 Sept 2012).
  • Santos, B. M., J. P. Morales-Payan, W. M. Stall, and T. A. Bewick. 1997a. Influence of tuber size and shoot removal on purple nutsedge (Cyperus rotundus) regrowth. Weed Science 45: 681–683. (Available online at: http://www.jstor.org/stable/4045894) (verified 10 Sept 2012).
  • Santos, B. M., J. P. Morales-Payan, W. M. Stall, T. A. Bewick, and D. G. Shilling. 1997b. Effects of shading on the growth of nutsedges (Cyperus spp.). Weed Science 45: 670–673. (Available online at: http://www.jstor.org/stable/4045892) (verified 10 Sept 2012).
  • Santos, B. M., J. P. Morales-Payan, W. M. Stall, and T. A. Bewick. 1998. Influence of purple nutsedge (Cyperus rotundus) density and nitrogen rate on radish (Raphanus sativus) yield. Weed Science 46: 661–664. (Available online at: http://www.jstor.org/stable/4045916) (verified 10 Sept 2012).
  • Shamsi, S. R. A., F. A. Al-ali, and S. M. Hussain. 1978. Temperature and light requirements for the sprouting of chilled and unchilled tubers of the purple nutsedge, Cyperus rotundus. Physiologia Plantarum 44: 193–196. (Available online at: http://dx.doi.org/10.1111/j.1399-3054.1978.tb08617.x) (verified 10 Sept 2012).
  • Wang, G., M. E. McGiffen, Jr., and E. J. Ogbuchiekwe. 2008. Crop rotation effects on Cyperus rotundus and C. esculentus population dynamics in southern California vegetable production. Weed Research 48: 420–428. (Available online at: http://dx.doi.org/10.1111/j.1365-3180.2008.00649.x) (verified 10 Sept 2012).
  • Webster, T. M. 2003. High temperatures and durations of exposure reduce nutsedge (Cyperus spp.) tuber viability. Weed Science 51: 1010–1015. (Available online at: http://dx.doi.org/10.1614/WS-03-018R) (verified 10 Sept 2012).
  • Webster, T. M. 2005a. Mulch type affects growth and tuber production of yellow nutsedge (Cyperus esculentus) and purple nutsedge (Cyperus rotundus). Weed Science 53: 834–838. (Available online at: http://dx.doi.org/10.1614/WS-05-029R.1) (verified 10 Sept 2012).
  • Webster, T. M. 2005b. Patch expansion of purple nutsedge (Cyperus rotundus) and yellow nutsedge (Cyperus exculentus) with and without polyethylene mulch. Weed Science 53: 839–845. (Available online at: http://dx.doi.org/10.1614/WS-05-045R.1) (verified 10 Sept 2012).
  • William, R. D. 1976. Purple nutsedge: Tropical scourge. HortScience 11: 357–364.
  • William, R. D., and G. F. Warren. 1975. Competition between purple nutsedge and vegetables. Weed Science 23: 317–323. (Available online at: http://www.jstor.org/stable/4042570) (verified 10 Sept 2012).

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 5134

Weed Profile: Pigweeds (Amaranthus spp.)

mer, 2012/09/12 - 16:12

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Abstract

Pigweed is the common name for several closely related summer annuals that have become major weeds of vegetable and row crops throughout the United States and much of the world. Most pigweeds are tall, erect-to-bushy plants with simple, oval- to diamond-shaped, alternate leaves, and dense inflorescences (flower clusters) comprised of many small, greenish flowers. They emerge, grow, flower, set seed, and die within the frost-free growing season.

Pigweeds thrive in hot weather, tolerate drought, respond to high levels of available nutrients, and are adapted to avoid shading through rapid stem elongation. They compete aggressively against warm season crops, and reproduce by prolific seed production.

In organic production systems, pigweeds can be managed through a combination of:

  • Timely cultivation, flame weeding, and manual removal
  • Stale seedbed
  • Mulching
  • Crop rotations that vary timing of tillage and other operations
  • Cover crops and competitive cash crops
  • Measures to prevent or minimize production of viable seeds
Introduction

Virtually every farmer in North America knows and grapples with pigweed, a term that covers several species in the genus Amaranthus, including:

  • redroot pigweed (A. retroflexus)
  • smooth pigweed (A. hybridus)
  • Powell amaranth (A. powelii)
  • Palmer amaranth (A. palmeri)
  • spiny amaranth (A. spinosus)
  • tumble pigweed (A. albus)
  • prostrate pigweed (A. blitoides)
  • waterhemp (A. tuberculatus = A. rudis)

These heat-loving summer annuals emerge after the spring frost date, grow rapidly, compete vigorously against warm-season crops, reproduce by seed, and die with the fall frost. Pigweeds are major weeds of warm season vegetables (Webster, 2006) and row crops (Sellers et al., 2003).

Also called amaranths, pigweeds are native to parts of North and Central America. Crop cultivation and human commerce have opened new niches, allowing pigweeds to invade agricultural ecosystems throughout the Americas, and parts of Europe, Asia, Africa, and Australia. Most amaranths make nutritious green vegetables or grain crops, and deliberate planting for food has helped some weedy species spread around the world. However, none of the pigweeds discussed here is grown commercially for grain, and modern grain amaranth varieties are not considered major agricultural weeds.

Pigweed problems have increased in no-till production systems with conventional herbicides, which leave weed seeds at the surface and select for herbicide-resistant populations (Sellers et al., 2003). However, high pigweed populations can occur on organic and non-organic farms, and in conventional, conservation, and no-till systems.

Description and Identification

Pigweeds are easy to recognize, yet correct identification of pigweed species can be tricky. Two or more pigweed species often occur together in the same field (Fig. 1), significant variation can occur within a species, and interspecific hybrids occasionally occur (Sellers et al., 2003). Some researchers consider tall waterhemp and common waterhemp a single species: A. tuberculatus (Pratt and Clark, 2001). Kansas State University Extension has published an excellent pigweed identification guide with photo illustrations and a key to distinguish mature plants of nine different weedy amaranths (Horak et al., 1994).

Palmer amaranth and smooth pigweed
Figure 1. Two pigweed species, tentatively identified as Palmer amaranth (left) and smooth pigweed (right), grow at the edge of a plastic mulched bed in organic vegetable production in Clemson, South Carolina. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Newly emerging pigweed seedlings open a pair of long, narrow cotyledons, about 0.5 inch long by 0.1 inch wide, followed by the first true leaves, which are broader in outline (Fig. 2). Plants form moderately deep, branching taproots, and may show a distinct reddish coloration on roots, lower stems, and undersides of leaves.

Pigweed seedlings
Figure 2. In this flush of summer annual weed seedlings, pigweed (Amaranthus sp.) can be distinguished by its pair of long, narrow cotyledons (seed leaves), and, on older seedlings, true leaves that are much more broadly oval in outline. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Most pigweeds grow into large, erect-to-bushy plants, 2–7 feet in height, with simple, petiolate (stalked) leaves arranged alternately (singly) on stems (Fig. 3a). Leaf blades are generally oval-to-diamond shaped, and 2–6 inches long. Prostrate pigweed forms a low, spreading mat, with smaller (about one inch) leaves that are distinctly notched at the tip (Fig. 3b).

Sooth pigweed; prostrate pigweed
Figure 3. a. These smooth pigweeds in early heading are about four feet tall. b. Prostrate pigweed forms a low, spreading mat. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Individual pigweed flowers are small, inconspicuous, and usually greenish in color. Male and female flowers are borne on the same plant (most species) or separate plants (waterhemp, Palmer amaranth). Each plant bears thousands of flowers in small clusters in leaf axils, or larger, often branched, densely-packed spikes at the tips of main stems and major branches (Fig. 4). Female flowers form single, small, round, usually shiny, dark reddish-brown-to-black seeds, roughly 0.04 inch in diameter (Fig. 5). About 50,000–90,000 seeds weigh one ounce.

Spiny amaranth and smooth pigweed in bloom
Figure 4. Spiny amaranth (left) and smooth pigweed (right) in bloom. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Seeds of tumble and redroot pigweed.
Figure 5. (a) Seeds of tumble pigweed. (b) Seeds of redroot pigweed, magnified, showing dark, shiny seed coat of mature seeds. Figure credits: (a) Steve Dewey, Utah State University, Bugwood.org. (b) Ken Chamberlain, Ohio State University, Bugwood.org.

See Table 1 below for a quick guide to eight common North American pigweed species, with links to additional information about each.

Table 1. Eight North American pigweed species at a glance.
Common and Scientific Name Growth Habit Inflorescence* Geographic Range** Other Plant Characteristics Redroot Pigweed Amaranthus retroflexus Erect, branched, 2–7 ft Stiff, branched terminal spikes, individual branches usually <2 in long, thicker than pencil Throughout North America including Alaska Upper stem and leaves usually covered with fine hairs; leaf blades large (6 in) on vigorous plants Smooth Pigweed Amaranthus hybridus Erect, branched, 2–7 ft Soft, highly branched terminal spikes, individual branches thinner than pencil Throughout North America Similar to redroot but highly variable, many local variants, may hybridize with closely related species Palmer Amaranth Amaranthus palmeri Erect, branched, 2–10 ft Long (to 18 in), simple or sparingly branched terminal spikes; male soft, female bristly Southern half of U.S., Great Plains, Mexico Extremely rapid, aggressive growth in hot climates, male and female flowers on separate plants; plants smooth and hairless Powell Amaranth Amaranthus powellii Erect, branched, 2–6 ft Stiff, branched terminal spikes, branches 4–8 in long, thicker than pencil, held close to main axis Throughout North
America First true leaves narrower and more tapered toward tip than redroot or smooth; plant may be smooth or hairy Spiny Amaranth Amaranthus spinosus Erect to bushy 1–4 ft Slender, branched terminal spikes mostly male flowers; axillary clusters mostly female Throughout North America, but mostly Southeastern U.S. Pair of stiff, sharp ½-in spines at base of each leaf; stems smooth, hairless, often red Waterhemp Amaranthus rudis or A. tuberculatus*** Erect, tall 3–10 ft Slender, simple or branched terminal spikes Throughout U.S. and southern Canada except driest areas Male and female flowers on separate plants; stems and leaves smooth and hairless; leaves often longer and narrower than other species Prostrate Pigweed Amaranthus blitoides Prostrate mat to 3 ft across Small, dense clusters in leaf axils Throughout U.S. and southern Canada Leaves small (blade about 1 in) with distinct notch at tip; seeds dull black, larger than in other pigweeds (0.06 in) Tumble Pigweed Amaranthus albus Globular bush, 1–3 ft diameter Small, dense clusters in leaf axils Throughout North America Mature plants break off at ground level, and are carried by wind, dispersing seeds; stems white to pale green, leaves light green * Small clusters of flowers are usually present in leaf axils of all amaranths
** Within North America (Canada, U.S., Mexico); many species have become naturalized on other continents.
*** Some authors recognize two species, common waterhemp (A. rudis) and tall waterhemp (A. tuberculatus); others consider them subspecies, or synonymous. Life Cycle, Reproduction, Seed Dispersal, Seed Dormancy, and Germination

Pigweeds are frost-tender summer annuals that emerge, grow, flower, and form mature seed within the frost-free period. Seedlings emerge over an extended period, with major flushes in late spring or early summer (Fig. 6). In most species, flowering and seed development take place mainly after the summer solstice, in response to shortening daylengths.

Smooth pigweed seedlings
Figure 6. A flush of smooth pigweed seedlings on a vegetable farm in the Tidewater region of Virginia, photographed on June 20, 2010, about two weeks after emergence. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Pigweeds reproduce entirely by seed. A single large plant can mature 100,000–600,000 seeds, and populations of 0.1–1 plants per square foot can shed 10,000–45,000 seeds per square foot, or 0.4–2 billion per acre (Massinga et al., 2001; Sellers et al., 2003). This prolific seed production makes pigweeds especially difficult to manage, since successful maturation of just one plant per 10,000 emerging seedlings can allow pigweed populations to increase severalfold from one year to the next.

Pigweeds typically begin to flower and shed pollen (anthesis) about six weeks after emergence (WAE), although flowers can occur as early as 3 WAE or as late as 9 WAE (Huang et al., 2000; Keeley, et al., 1987; Shrestha and Swanton, 2007). Flowers open about 1–2 weeks after flower buds first become visible to the unaided eye.

Reported time intervals from pollination to formation of viable seeds range from 7–12 days in waterhemp (Bell and Tranel , 2010) to 6 weeks in field populations of redroot pigweed in Ontario (Shrestha and Swanton, 2007). In California, Palmer amaranth formed viable seeds 2–6 weeks after flowering (Keeley et al., 1987). Seeds become viable at about the same time that they develop their mature dark brown or black color.

Reproductive development is accelerated by shortening daylength after the summer solstice in field populations (Keeley et al., 1987), and proceeds faster in short (~12 hour) than in longer (≥14 hour) photoperiods in a growth chamber (Huang et al., 2000). Although most seed production occurs in late summer and early fall, some mature seeds have been found in smooth pigweed seed heads at the summer solstice in Virginia (personal observation).

The ability of pigweed plants uprooted or severed at flowering to complete seed maturation has not been researched. However, in upstate New York, 2–4-inch fragments of Powell amaranth inflorescences lying on the soil surface were found to contain black seeds 3 weeks after the weeds were disked down at flowering (Charles Mohler, Cornell University, pers. commun.). Apparently, if pollination takes place before pigweeds are pulled or chopped, some potential exists for viable seed production.

Pigweed seeds are dispersed to new locations by irrigation or flood water, manure, and soil clinging to footwear, tractor tires, or tillage tools. In addition, tumble pigweed actively disperses seeds when mature plants break off and move with the wind.

Pigweed seeds have multiple dormancy mechanisms, so that seeds produced in a given season germinate at different times over the next several years, thereby enhancing the weed's long-term persistence (Egley, 1986). Newly shed pigweed seeds are mostly dormant, and become less so by the following spring. Germination is promoted by high temperatures (95 °F), diurnally fluctuating temperatures (e.g., 85–95 °F day, ~ 70 °F night), and sometimes light (Guo and Al-Khatib, 2003; Schonbeck and Egley, 1980 and 1981 Steckel et al., 2004).

Pigweed emerges most readily from the top 0.5–1.0 inch of the soil profile, with few emerging from seeds located deeper than one inch (Mohler and Di Tommaso, unpublished). The seeds require adequate moisture and good seed–soil contact to absorb moisture and germinate. More deeply buried seeds remain dormant and viable for several years, and germinate when brought to the surface by tillage or cultivation. Although flushes of emergence commonly follow seedbed preparation or cultivation, increasing pigweed problems in agronomic crops have been attributed to widespread adoption of no-till and minimum-tillage, which leave recently-shed weed seeds at or near the soil surface (Sellers et al., 2003).

Growth Habit and Impact on Crops

Pigweeds have the C4 photosynthetic pathway, which confers an ability to grow rapidly at high temperatures and high light levels, to tolerate drought, and to compete aggressively with warm-season vegetables for light, moisture, and nutrients. Growth is related to cumulative Growing Degree Days, with a base temperature of 50 °F (Shrestha and Swanton, 2007; Horak and Loughin, 2000); thus, pigweeds grow much faster in hot climates than in northern regions with cooler summers.

Erect pigweed species can rapidly overtop short crops like broccoli or snap bean. In taller crops like corn, pigweeds respond to canopy shade by increasing stem growth and deploying leaves higher on the plant, thereby intercepting a larger fraction of available light (Massinga et al., 2003; McLachlan et al, 1993). One to three pigweed plants per 10 feet of row emerging with corn or soybean can cause significant yield losses (Klingman and Oliver, 1994; Knezevic et al., 1994; Massinga et al., 2001) Pigweeds that emerge several weeks after the crop has emerged exert much less effect on yields.

Pigweeds are highly responsive to nutrients, especially the nitrate form of nitrogen (N) (Blackshaw and Brandt, 2008; Teyker et al., 1991). Fertilization enhances both weed biomass and seed production. In addition, nitrate can stimulate pigweed seed germination (Egley, 1986). A mulch of legume cover crop residues has been observed to enhance pigweed emergence in some years (Fig. 7), likely as a result of rapid mineralization of legume N (Teasdale and Mohler, 2000).

Pigweed response to different cover crops
Figure 7. In this field trial, a flush of pigweed competes against broccoli planted no-till into killed hairy vetch (foreground), while broccoli planted in killed rye or rye–vetch are relatively free of pigweed (background). Rapid N mineralization from the all-legume cover crop residues apparently stimulated pigweed germination and growth. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Because the small seeds have minimal nutrient reserves, pigweed seedlings are initially more dependent on readily available nutrients from the soil, especially phosphorus (P) and potassium (K), than larger-seeded plants such as corn, beans, and cucurbits (Hoveland et al., 1976; Mohler, 1996). However, in studies conducted on organic (muck) soils in Florida, smooth pigweed and spiny amaranth were less responsive than lettuce to P levels, and a band application of P fertilizer improved the crop's ability to compete against these weeds (Santos et al., 1997; Shrefler et al., 1994).

Pigweeds are shade intolerant, and the growth and reproduction of individuals that emerge under a heavy crop canopy are substantially reduced. However, rapid stem elongation allows pigweeds to escape shading in many cropping situations. Late-season pigweeds that break through established cucurbit, tomato, pepper, and other vegetables can promote crop disease by reducing air circulation, interfere with harvest, and set many thousands of seeds (Fig. 8).

Pigweed heading in squash
Figure 8. The pigweed emerged several weeks after squash planting and did not affect yield. By the end of crop harvest, however, each weed matured thousands of seeds, and will make a heavy deposit into the weed seed bank unless they are removed promptly. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Pigweeds are reported to host pest nematodes (Meloidogyne spp.) and many vegetable crop pathogens, including fungi that cause early blight in potato and tomato (Alternaria solani), lettuce drop (Sclerotinia sclerotiorum) and southern blight (Sclerotium rolfsii) in a wide range of crops. Viral pathogens such as cucumber mosaic virus and tomato spotted wilt virus can also be transmitted from pigweeds (Mohler and DiTommaso, unpublished).

Pigweeds have become the focus of biocontrol efforts with fungal pathogens and plant-feeding insects, although no biocontrol products have yet become available to farmers. The amaranth flea beetle (Disonycha glabrata) occurs throughout much of the United States (Tisler, 1990), feeds on pigweed foliage (Fig. 9), and may become a significant natural enemy of pigweed in some areas, including Floyd County, Virginia (personal observation). However, it usually does not control the weeds, and occasionally feeds on some vegetable seedlings.

Amaranth flea beetle
Figure 9. The amaranth flea beetle feeds on pigweed foliage, and has been observed to cause substantial defoliation and reduce weed vigor in some parts of Virginia. This insect can occasionally become a pest in beet and chard by feeding on seedlings. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Management

Organic farmers manage pigweeds by taking advantage of their points of vulnerability. The small seeds have minimal nutrient reserves; thus seedlings can emerge only from seeds located within an inch of the soil surface, and are immediately dependent on the soil for readily available nutrients. Transplanted and large-seeded crops have substantial nutrient reserves, and can gain a competitive edge over pigweed seedlings if slow-release nutrient sources are used.

The delicate seedlings are readily killed by severing, uprooting, burial, or heat. Timely flame weeding or cultivation with any of a variety of implements can knock out a flush of pigweed seedlings. Emerging pigweed is also susceptible to shading and physical hindrance by mulch. A field study at Beltsville, Maryland documents the greater sensitivity of pigweed to suppression with organic mulches relative to several other common weeds: redroot pigweed > lamb's quarter > giant foxtail > velvetleaf (Teasdale and Mohler, 2000).

Timely action is vital, as pigweeds rapidly become harder to kill once they grow taller than one inch and develop four or more true leaves (Fig. 10). In cool climates, pigweed seedlings may remain vulnerable to cultivation for up to 4 WAE (Weaver and McWilliams, 1980); however in warmer climates, they can grow to 2–4 inches within 2 WAE (Sellers et al., 2003).

Pigweed seedlings
Figure 10. The pigweed seedling on the right is at the vulnerable stage, at which it can be readily killed by shallow cultivation or flaming, or blocked by mulch. When pigweed grows as large as the seedling on the left, it becomes more difficult to kill, requiring more vigorous cultivation. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Pigweed populations readily adapt to production systems and control tactics. For example, seed germination responses show adaptive changes to different crop rotations (Brainard et al., 2007), and widespread herbicide resistance has been reported in several species (Fugate, 2009; Volenberg et al., 2007). Thus, reliance on a single management tool or the same strategy year after year will likely yield diminishing returns over time.

When used in combination, the practices described below can provide effective management of pigweed in organic systems.

Cultivation and Flame Weeding

Monitor crops regularly for weed emergence. Cultivate when pigweeds are in the cotyledon stage, or before they reach one inch in height, working as close to the crop row as practical. When the crop is sufficiently established, set cultivators to move an inch or so of soil into rows to bury small weeds. The surface layer of loose, dry soil left by cultivation (dust mulch) deters additional pigweed germination. Avoid recompacting the soil, as compaction can promote another flush of emergence (Fig. 11).

Dust mulch from cultivation suppresses weed seed germination
Figure 11 Cultivation left a dust mulch around these young squash plants, thereby discouraging germination of pigweed and other small-seeded weeds. However, foot traffic recompacted the soil enough to re-establish seed–soil contact near the surface, thereby allowing weed seeds to imbibe moisture, germinate, and grow in the footprints. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Flame weeding can remove pigweed and other broadleaf seedlings just before crop emergence. It is often used for slow-starting crops like carrot, beet, and parsnip. Because flaming usually does not kill grass weed seedlings, it is not recommended where grasses make up a significant portion of the weed flora.

Mowing and Grazing

Once the crop is too large to cultivate by tractor, farmers often mow, cut, or pull weeds in alleys to maintain air circulation around the crop, facilitate harvest, and prevent weed propagation. This should be done before pigweed flowers open (within a few days after flower heads first become visible) to prevent viable seed formation.

Some farmers mow alleys between wide rows or plastic mulched beds with a push mower or line trimmer as a soil-saving alternative to cultivation. Two timely mowings prior to canopy closure have given adequate between-row control of giant foxtail, pigweeds, and ragweed in soybean planted in a 30-inch row spacing (Donald, 2000).

Most pigweeds are highly palatable to livestock. However, mature seeds pass through the animals' digestive tracts unharmed, and manure is a notorious source of pigweed seeds. Thus, pigweed should be grazed while still vegetative. Note also that the National Organic Program requires a 120-day interval between manure deposits by grazing animals and the next food-crop harvest.

Mulching

Mulching can be an effective control tactic for pigweeds in vegetable production. An organic mulch, such as 3–4 inches of straw or hay (~5–10 tons/ac), applied within a day after cultivating an established crop, can reduce subsequent pigweed emergence by 90%. Alternatively, a synthetic mulch such as black plastic can be laid before crop planting, and alley weeds controlled by cultivation, mowing, organic mulch, or cover crop. Note: If plastic or other synthetic mulch is used to organic crops, it must be removed from the field at the end of the harvest or growing season.

Organic no-till transplanting of tomato and other summer vegetables into roll–crimped or mowed winter cover crops can control light-to-moderate pigweed populations. Rye residues release natural plant growth inhibitors (allelochemicals) that suppress pigweed and some other annual weeds (Barnes and Putnam, 1983; Putnam et al., 1983) without affecting transplanted vegetables.

Nutrient and Moisture Management

Use slow-release sources of N and other crop nutrients, and avoid broadcast application of faster-release materials like blood meal and bone meal, which can give pigweed the jump on the crop. For heavy feeders like broccoli or spinach that need some quick N, band or side dress materials within or near the crop row at the onset of rapid crop growth.

Use in-row drip irrigation to provide water and liquid organic fertilizer directly to the crop without feeding and watering between-row weeds. Subsurface drip lines can provide moisture to the crop and leave the soil surface dry, thereby minimizing within-row weed emergence.

Crop Rotation, Planting Schedules, and Stale Seedbed

Plan crop rotation and schedule field operations to disrupt pigweed life cycles. Avoid providing an open niche (bare soil) year after year for pigweed emergence in late spring to early summer. Alternate warm- and cool-season vegetables. Consider delaying seedbed preparation for a summer vegetable until after the time of peak pigweed emergence. After several years of intensive vegetable production, rotate the field to perennial sod (e.g., orchardgrass–red clover) for two or three years to disrupt pigweed life cycles and encourage weed seed predation.

If pigweed populations are high (Fig. 12), prepare a stale seedbed in late spring to draw down the weed seed bank. Till or cultivate, then roll or cultipack the soil to improve seed–soil contact, thereby promoting weed germination. Sprinkle irrigate if the soil is dry. Repeat cultivation as needed. Just before crop planting or crop emergence, use shallow cultivation and leave the surface loose to discourage additional weed germination. The final flush can also be killed by flame if grass weeds are few or absent.

Spiny amaranth seedlings
Figure 12. A carpet of spiny amaranth seedlings arises from a large weed seed bank. A stale seedbed or cultivated fallow is needed to bring this situation under control. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Crop Competition and Cover Cropping

With good early-season weed control, vigorous crops like tomato, sweet potato, and winter squash can tolerate later-emerging pigweed. However, crop competition may not control pigweeds, because of their shade-avoidance response and ability to break through the crop canopy through rapid stem elongation.

Competitive summer cover crops such as buckwheat, sorghum–sudangrass, cowpea, and forage soybean are often used to suppress weeds between spring and fall vegetable crops. In Florida, cowpea, sunnhemp, or velvetbean cover crops seeded at high rates reduced but did not eliminate smooth pigweed growth (Collins et al., 2008).

When using summer cover crops to combat pigweeds, seed at high rates (1.5–2 times normal), and use good seeding methods to obtain a weed-suppressive cover crop stand. Combine cowpea, forage soybean, or other summer legume with a tall grass like pearl millet or sorghum–sudangrass to develop a canopy that is both tall and dense. Watch the crop closely; if a significant amount of pigweed grows with it, terminate the crop promptly when weed flower heads first appear.

Managing the Pigweed Seed Bank

Because pigweeds produce seed so prolifically, it is critical to minimize the annual seed rain onto the soil. Although stringent weed control for six years can reduce the pigweed seed bank by 99%, relaxing weed control allows seed numbers to recover to near original levels within three years (Schweizer and Zimdahl, 1984). Pigweed that emerges after a crop's minimum weed-free period may not reduce the crop's yield, but it should be pulled or cut before flowering to prevent formation of mature seeds.

It may pay to walk fields of maturing crops to pull or chop out large weeds; small, stunted pigweeds below a crop canopy form only small numbers of seeds. If flower heads are already formed, remove severed or uprooted pigweed plants from the field. If pigweed plants have already formed seed, note that many of the seeds will stay in the head until winter. Therefore, removing the weeds in early fall can still significantly reduce the pigweed seed rain.

In the event that a heavy pigweed seed rain occurs, some weed scientists recommend inversion tillage to move seeds to a depth from which they cannot emerge (Mohler and Di Tommaso, unpublished). Although 5–14% of redroot pigweed and waterhemp seeds have survived 9–12 years burial at 8-inch depth in Nebraska (Burnside et al., 1996), others have reported that pigweed seeds are fairly short lived (3–4 years) in the soil in more humid regions such as Mississippi and Illinois (Buhler and Hartzler, 2001; Egley and Williams, 1990; Steckel et al., 2007). Moldboard plowing has been reported to increase pigweed emergence if weed populations are low, but to decrease emergence if populations are high as a result of a recent seed rain (Schweizer and Zimdahl, 1984).

When using inversion tillage to manage a heavy seed deposit, moldboard plow the field once, then avoid deep tillage for the next several years to allow buried seeds to lose viability.

This article is part of a series discussing the invasive family of Pigweeds. For more information, see the following articles: References Cited
  • Barnes, J. P., and A. R. Putnam. 1983. Rye residues contribute weed suppression in no-tillage cropping systems. Journal of Chemical Ecology 9: 1045–1057. (Available online at: http://dx.doi.org/10.1007/BF00982210) (verified 10 Sept 2012).
  • Bell, M. S., and P. J. Tranel. Time requirement from pollination to seed maturity in waterhemp (Amaranthus tuberculatus). Weed Science 58: 167–173. (Available online at: http://dx.doi.org/10.1614/WS-D-09-00049.1) (verified 10 Sept 2012).
  • Blackshaw, R. E., and R. N. Brandt. 2008. Nitrogen fertilizer rate effects on weed competitiveness is species dependent. Weed Science 56: 743–747. (Available online at: http://dx.doi.org/10.1614/WS-08-065.1) (verified 10 Sept 2012).
  • Brainard, D. C., A. DiTommaso, and C. A. Mohler. 2007. Intraspecific variation in seed characteristics of Powell amaranth (Amaranthus powellii) from habitats with contrasting crop rotation histories. Weed Science 55: 218–226. (Available online at: http://dx.doi.org/10.1614/WS-06-134.1) (verified 10 Sept 2012).
  • Buhler, D. D. and R. G. Hartzler. 2001. Emergence and persistence of seed of velvetleaf, common waterhemp, woolly cupgrass, and giant foxtail. Weed Science 49: 230–235. (Available online at: http://dx.doi.org/10.1614/0043-1745(2001)049%5B0230:EAPOSO%5D2.0.CO;2) (verified 10 Sept 2012).
  • Burnside, O. C., R. G. Wilson, S. Weisberg, and K. G. Hubbard. 1996. Seed longevity of 41 weed species buried 17 years in eastern and western Nebraska. Weed Science 44: 74–86. (Available online at: http://www.jstor.org/stable/4045786) (verified 10 Sept 2012).
  • Collins, A. S., C. A. Chase, W. M. Stall, and C. M. Hutchinson. 2008. Optimum densities of three leguminous cover crops for suppression of smooth pigweed (Amaranthus hybridus). Weed Science 56: 753–761. (Available online at: http://dx.doi.org/10.1614/WS-07-101.1) (verified 10 Sept 2012).
  • Donald, W. W. 2000. Between-row mowing + in-row band-applied herbicide for weed control in Glycine max. Weed Science 48: 487–500. (Available online at: http://www.jstor.org/stable/4046280) (verified 10 Sept 2012).
  • Egley, G. H. 1986. Stimulation of weed seed germination in soil. Reviews of Weed Science 2: 67–89..
  • Egley, G. H., and R. D. Williams. 1990. Decline of weed seeds and seedling emergence over five years as affected by soil disturbances. Weed Science 38: 504–510. (Available online at: http://www.jstor.org/stable/4045064) (verified 10 Sept 2012).
  • Fugate, L. 2009. Pigweed causing farmers to rethink farming methods. University of Arkansas Division of Agriculture Cooperative Extension Service News - October 2009.
  • Guo, P., and K. Al-Khatib. 2003. Temperature effects on germination and growth of redroot pigweed (Amaranthus retroflexus), Palmer amaranth (A. palmeri), and common waterhemp (A. rudis). Weed Science 51: 869–875. (Available online at: http://dx.doi.org/10.1614/P2002-127) (verified 10 Sept 2012).
  • Horak, M. J., and T. M. Loughin. 2000. Growth analysis of four Amaranthus species. Weed Science 48: 347–355. (Available online at: http://dx.doi.org/10.1614/0043-1745(2000)048%5B0347:GAOFAS%5D2.0.CO;2) (verified 10 Sept 2012).
  • Horak, M. J., D. E. Peterson, D. J. Chessman, and L. M. Wax. 1994. Pigweed identification: a Pictoral guide to the common pigweeds of the Great Plains. 12 pp. (Available online at: http://www.ksre.ksu.edu/library/crpsl2/s80.pdf) (verified ).
  • Hoveland, C. S., G. A. Buchanan, and M. C. Harris. 1976. Response of weeds to soil phosphorus and potassium. Weed Science 24: 194–201. (Available online at: http://www.jstor.org/stable/4042586) (verified 10 Sept 2012).
  • Huang, J. Z., A. Shrestha, M. Tollenar, W. Deen, H. Rahimian, and C. J. Swanton. 2000. Effect of photoperiod on the phenological development of redroot pigweed (Amaranthus retroflexus L.). Canadian Journal of Plant Science 80: 929–938.
  • Keeley, P. E., C. H. Carter, and R. J. Thullen. 1987. Influence of planting date on growth of Palmer amaranth (Amaranthus palmeri). Weed Science 35: 199–204. (Available online at: http://www.jstor.org/stable/4044391) (verified 10 Sept 2012).
  • Klingman, T. E., and L. R. Oliver. 1994. Palmer amaranth (Amaranthus palmeri) interference in soybeans (Glycine max). Weed Science 42: 523–527. (Available online at: http://www.jstor.org/stable/4045448) (verified 10 Sept 2012).
  • Knezevic, S. Z., S. F. Weise, and C. J. Swanton. 1994. Interference of redroot pigweed (Amaranthus retroflexus) in corn (Zea mays). Weed Science 42: 568–573. (Available online at: http://www.jstor.org/stable/4045456) (verified 10 Sept 2012).
  • Massinga, R. A., R. S. Currie, M. J. Horak, and J. Boyer, Jr. 2001. Interference of Palmer amaranth in corn. Weed Science 49: 202–208. (Available online at: http://dx.doi.org/10.1614/0043-1745(2001)049%5B0202:IOPAIC%5D2.0.CO;2) (verified 10 Sept 2012).
  • Massinga, R. A., R. S. Currie, and T. P. Trooien. 2003. Water use and light interception under Palmer amaranth (Amaranthus palmeri) and corn competition. Weed Science 51: 523–531. (Available online at: http://dx.doi.org/10.1614/0043-1745(2003)051%5B0523:WUALIU%5D2.0.CO;2) (verified 10 Sept 2012).
  • McLachlan, S. M., M. Tollenaar, C. J. Swanton, and S. F. Weise. 1993. Effect of corn-induced shading on dry matter accumulation, distribution, and architecture of redroot pigweed (Amaranthus retroflexus). Weed Science 41: 568–573. (Available online at: http://www.jstor.org/stable/4045424) (verified 10 Sept 2012).
  • Mohler, C. A. 1996. Ecological bases for the cultural control of annual weeds. Journal of Production Agriculture 9:  468–474..
  • Mohler, C. A., and A. DiTommaso. Unpublished. Manage Weeds on Your Farm: a Guide to Ecological Strategies. Department of Crop and Soil Sciences, Cornell University. Pre-publication draft, version 5.1. Publication anticipated in 2012.
  • Pratt, D. B., and L. G. Clark. 2001. Amaranthus rudis and A. tuberculatus—one species or two? Journal of the Torrey Botanical Society 128: 282–296. (Available online at: http://www.jstor.org/stable/3088718) (verified 10 Sept 2012).
  • Putnam, A. R., J. DeFrank, and J. P. Barnes. 1983. Exploitation of allelopathy for weed control in annual and perennial cropping systems. Journal of Chemical Ecology 9: 1001–1010. (Available online at: http://dx.doi.org/10.1007/BF00982207) (verified 10 Sept 2012).
  • Santos, B. M., J. A. Dusky, D. G. shilling, W. M. Stall, and T. A. Bewick. 1997. Effect of phosphorus fertility on competitive interactions of smooth pigweed (Amaranthus hubridus), spiny amaranth (Amaranthus spinosus), and common purslane (Portulaca oleracea) with lettuce. Weed Science Society of America Abstracts 37: 54.
  • Schonbeck, M. W., and G. H. Egley. 1980 Redroot pigweed (Amaranthus retroflexus) seed germination responses to afterripening, temperature, ethylene, and some other environmental factors. Weed Science 28: 543–548. (Available online at: http://www.jstor.org/stable/4043277) (verified 10 Sept 2012).
  • Schonbeck, M. W., and G. H. Egley. 1981. Changes in sensitivity of Amaranthus retroflexus L. seeds to ethylene during preincubation. II. Effects of alternating temperature and burial in soil. Plant, Cell, and Environment 4: 237–242. (Available online at: http://dx.doi.org/10.1111/1365-3040.ep11611005) (verified 10 Sept 2012).
  • Schweizer, E. E., and R. L. Zimdahl. 1984. Weed seed decline in irrigated soil after six years of continuous corn (Zea mays) and herbicides. Weed Science 32: 76–83. (Available online at: http://www.jstor.org/stable/4043886) (verified 10 Sept 2012).
  • Sellers, B. A., R. J. Smeda, W. G. Johnson, J. A. Kendig, and M. R. Ellersieck. 2003. Comparative growth of six Amaranthus species in Missouri. Weed Science 51: 329–333. (Available online at: http://dx.doi.org/10.1614/0043-1745(2003)051%5B0329:CGOSAS%5D2.0.CO;2) (verified 10 Sept 2012).
  • Shrefler, J. W., J. A. Dusky, D. G. Shilling, B. J. Brecke, and C. A. Sanchez. 1994. Effect of phosphorus fertility on competition between lettuce (Lactuca sativa) and spiny amaranth (Amaranthus spinosus). Weed Science 42: 556–560. (Available online at: http://www.jstor.org/stable/4045454) (verified 10 Sept 2012).
  • Shrestha, A., and C. J. Swanton. 2007. Parameterization of the phenological development of select annual weeds under noncropped field conditions. Weed Science 55: 446–454. (Available online at: http://dx.doi.org/10.1614/WS-06-176.1) (verified 10 Sept 2012).
  • Steckel, L. E., C. L. Sprague, E. W. Stoller, and L. M. Wax. 2004. Temperature effects on germination of nine Amaranthus species. Weed Science 52: 217–221. (Available online at: http://dx.doi.org/10.1614/WS-03-012R) (verified 10 Sept 2012).
  • Steckel, L. E., C. L. Sprague, E. W. Stoller, L. M. Wax, and F. W. Simmons. 2007. Tillage, cropping system, and soil depth effects on common waterhemp (Amaranthus rudis) seed-bank persistence. Weed Science 55: 235–239. (Available online at: http://dx.doi.org/10.1614/WS-06-198) (verified 10 Sept 2012).
  • Teasdale, J. R., and C. L. Mohler. 2000. The quantitative relationship between weed emergence and the physical properties of mulches. Weed Science 48: 385–392. (Available online at: http://dx.doi.org/10.1614/0043-1745(2000)048%5B0385:TQRBWE%5D2.0.CO;2) (verified 10 Sept 2012).
  • Teyker, R. H., H. D. Hoelzer, and R. A. Liebl. 1991. Maize and pigweed response to nitrogen supply and form. Plant and Soil 135: 287–292. (Available online at: http://dx.doi.org/10.1007/BF00010918) (verified 10 Sept 2012).
  • Tisler, A. M. 1990. Feeding in the pigweed flea beetle, Disonycha glabrata Fab. (Coleoptera: Chrysomelidae), on Amaranthus retroflexus. Virginia Journal of Science 41: 243–245. (Available online at: http://www.vacadsci.org/vjsArchives/v41/41-3/41-243.pdf) (verified 10 Sept 2012).
  • Volenberg, D. S., W. L. Patzoldt, A. G. Hager, and P. J. Tranel. 2007. Responses of contemporary and historical waterhemp (Amaranthus tuberculatus) accessions to glyphosate. Weed Science 55: 327–333. (Available online at: http://dx.doi.org/10.1614/WS-06-121) (verified 10 Sept 2012).
  • Weaver, S. E., and E. L. McWilliams. 1980. The biology of Canadian weeds. 44. Amaranthus retroflexus L, A. powellii S. Wats and A. hybridus L. Canadian Journal of Plant Science 60: 1215–1234.
  • Webster, T. M. 2006. Weed survey – southern states. Vegetable, fruit and nut crops subsection. Proceedings of the Southern Weed Science Society 59: 260–277. (Available online at: http://www.swss.ws/NewWebDesign/Publications/Weed%20Survey%20Archives/Southern%20Weed%20Survey%202006%20Vegetables%20and%20Fruits.pdf) (verified 10 Sept 2012).

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 5120

Organic Vegetable Production Systems, Control Practices in Organic Weed Management

mer, 2012/09/12 - 16:08

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T879,934

Organic Vegetable Production Systems, Production of Specific Vegetable Crops

mer, 2012/09/12 - 16:08

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T879,891

Control Practices in Organic Weed Management

mer, 2012/09/12 - 16:08

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T934

Weed Management Strategies for Organic Tomato, Pepper, and Eggplant in the Southern United States

mer, 2012/09/12 - 15:59

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Abstract

In organic production, tomato, pepper, and eggplant are normally started indoors and transplanted to the field to give them a head start on the weeds. Crops kept free of weeds for the first 4–8 weeks (tomato) or 8–10 weeks (eggplant, pepper) after transplanting can usually outcompete later-emerging weeds. However, late-season weeds can interfere with harvest, promote disease, and propagate themselves.

The most serious weeds of solanaceous (nightshade family) vegetables in the southern United States include yellow and purple nutsedges, morning glories, pigweeds, solanaceous weeds that host tomato or pepper diseases, and several winter annual weeds that harbor tomato spotted wilt virus and an insect pest (thrips) that transmits the virus from weed to crop. Fields heavily infested with these weeds should not be planted in tomato, pepper, or eggplant, and should be rotated to weed-smothering cover crops or perennial sod until weed populations decline to tolerable levels.

Important weed-preventive measures include: crop rotations that alternate warm- and cool-season vegetables, cover cropping, providing sufficient but not excessive nutrients from slow-release organic sources, and stale seedbed (planting delayed after seedbed preparation to allow removal of the inital flush of weeds) to draw down the weed seed bank.

Most organic farmers use one of two strategies to control weeds in solanaceous vegetables:

  • Black plastic mulch laid just before transplanting; alley weeds managed by cultivation, mowing, or organic mulch.
  • Cultivation to remove early-season flushes of weeds, followed by hay, straw, or other organic mulch.

Variations on these strategies include opaque white or light-reflecting (aluminized) plastic mulch for later plantings when soil warming is not desired; spreading organic mulch over plastic films several weeks after planting to prevent excessive soil heating; and in situ organic mulch from mowed or roll-crimped cover crops.

This article outlines organic weed management strategies and techniques for tomato and related vegetable crops, based on an understanding of:

  • General principles of ecological weed management.
  • Crop growth habits, life cycles, cultural practices, and organic production systems.
  • Inherent vulnerabilities and strengths of tomato, pepper, and eggplant in relation to weeds.
Introduction

Tomato, pepper, and eggplant require good early-season weed control for successful production. Once established, these warm-season solanaceous vegetables become less vulnerable to competition from newly-emerging weeds, and tall tomato varieties can hinder weed growth by shading. late-season weeds must be managed to maintain adequate air circulation around crop foliage, and to prevent weed propagation during the long harvest season.

Crop Life Cycle and Growth Habit, Impacts of Weeds, and Minimum Weed-Free Period

Tomato, pepper, and eggplant are frost-tender summer annual crops in the solanaceous (nightshade) plant family. They require warm temperatures and an adequate supply of nutrients throughout the season to support good growth and yields. Newly-emerged seedlings are highly vulnerable to weed competition, and are usually started indoors in a weed-free potting mix. Once established, these crops become more tolerant of weed pressure. In tomato, the minimum weed-free period (to prevent yield losses to weed competition) has been estimated as the first 4–8 weeks after transplanting (Monks, 1993; Riggs et al., 1991; Teasdale and Colacicco, 1985; Weaver and Tan, 1983). Minimum weed-free periods for pepper and eggplant may be a little longer (8–10 weeks), owing to their slower development and shorter stature.

Because harvests continue until plants are killed by fall frost or disease, mature solanaceous crops remain in the field for a long period, during which additional weed management may be needed. Weeds that emerge after the crop's minimum weed-free period and are allowed to grow can interfere with harvest, promote disease by harboring pathogens or curtailing air circulation, or set seed.

When solanaceous vegetables reach a height of 12 inches or so, they tolerate some hilling up during cultivation, which buries and kills small within-row weeds. Hilling can benefit tomato by stimulating adventitious rooting from the stem, and diverting excess moisture away from the base of the plants (Diver et al., 2007). However, hilling can promote development of southern stem blight (Sclerotium rolfsii), and should be avoided where this pathogen is present (Louws, 2009).

Sweet and hot peppers, eggplant, and compact determinate varieties of tomato (varieties that grow to a fixed mature size and ripen all their fruit in a short period) form upright, bushy plants two to three feet tall. Indeterminate and semi-determinate tomato varieties form longer vines (five feet or more), and are normally staked or trellised.

Tomato prefers moderately warm conditions, giving best growth and production at daily mean air temperatures of 70–75 °F (Peet, 1996, Swaider et. al, 1992) and soil temperatures of 68–86 °F (Tindall et al., 1990). On good soil without hardpan (compact layer that restricts root growth), tomato forms a deep (five feet) root system, which can make established plants less susceptible to weed competition for soil moisture. The heavy foliage of a vigorous tomato crop can shade out weeds; however managing the crop to promote canopy closure can aggravate disease problems, leading to defoliation and yield losses.

Peppers require similar growing conditions to tomatoes, and prefer slightly warmer temperatures, especially the pungent varieties (Peet, 1996, Swaider et al., 1992). Unlike tomato, pepper has a shallow root system, which makes it more vulnerable to weed competition for soil moisture, and to detrimental root pruning from cultivation. Pepper is less prone to foliar diseases than tomato, and can be managed for canopy closure within the bed when disease pressure is light.

Eggplants prefer higher temperatures (daily means about 80 °F, nights not cooler than 65 °F) and a longer growing season than other solanaceous vegetables (Swaider et al., 1992). Like pepper, it can be managed to close canopy within the bed for suppression of late-emerging weeds. The crop develops a moderately deep root system (four feet), and can tolerate shallow cultivation for weed control.

Cultural Practices and Weed Management

Tomato, pepper, and eggplant are normally started in the greenhouse in a weed-free potting mix, and transplanted to the field at an age of 5–7 weeks (tomato) or 8–10 weeks (pepper, eggplant). Although some large scale conventional growers direct-seed solanaceous crops and apply selective herbicides, nearly all organic farmers minimize early-season weed competition by transplanting vigorous starts (Fig. 1). Tomato is set out after the last spring frost date; pepper and eggplant are often planted a few weeks later, when the soil is thoroughly warm.

Some farmers do succession plantings, with early crops set out in high tunnels before the last spring frost date, and field plantings continuing into late June or early July. Later plantings set out after the late spring flush of weed emergence allow the farmer to reduce weed pressure by preparing a stale seedbed or growing a weed-suppressive spring cover crop. When using the stale seedbed technique (also known as false seedbed), the farmer delays crop planting for several weeks after initial seedbed preparation to allow one or more flushes of weed seedlings to emerge. These are removed by shallow cultivation or flame weeding, and the crop is planted immediately after the final cultivation or flaming.

Vigorous tomato and pepper starts
Figure 1. (a) Vigorous tomato starts at 6 weeks after seeding, ready for transplanting. (b) Pepper starts 8 weeks after seeding. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Tomato, pepper, and eggplant require a moderate amount of available N, about 70–130 lb/ac from planting through early fruit set, and ample phosphorus (P) and potassium (K) to support good root growth and fruit quality, respectively (Swaider et al., 1992). Good organic soil management and attention to maintaining optimum growing conditions can enhance the crop's competitiveness toward weeds. Planting when temperatures are below optimum can prolong the minimum weed-free period. Providing too much available N early in the season can give weeds a competitive advantage over the crop, and excessive N later in the season may result in plants with dense foliage and few fruit.

Tomato is usually planted in single rows spaced 5–6 feet apart, either in raised beds or on the flat, with individual plants set 12–48 inches apart, depending on cultivar and production system. Some organic growers use wider row spacing (8–10 feet) to enhance air circulation, sometimes planting a low-growing vegetable or cover crop in the intervening spaces. Plants are normally supported (staked, trellised, or caged), and pruned (suckered) regularly to promote rapid drying of foliage for disease control, and to enhance fruit yield, quality, and ripening. Trellising and stake-and-weave systems that train the crop to a single line allow close cultivation or mulching under the plants, while cages limit weed control options near plant bases. Short varieties and processing tomatoes are sometimes grown without support (ground culture); however, the sprawling plants are difficult to keep weeded unless a plastic mulch is used.

Pepper and eggplant are often planted in staggered double rows, with plants spaced 18–24 inches apart in the row, and double rows on 5–7 foot centers. With this planting pattern, the crop often closes canopy within the bed, thereby suppressing the growth of late-emerging weeds. Pepper is sometimes staked to prevent the shallow-rooted plants from falling over during heavy fruit set.

Solanaceous crops respond very well to mulching. Many commercial organic producers routinely use synthetic mulch—most often black polyethylene film (black plastic)—for tomato, pepper, and eggplant (Fig. 2). Black plastic effectively controls most weeds and warms the soil, thereby promoting crop earliness and sometimes total yield. Drip irrigation is usually laid under the mulch to deliver water and liquid organic fertilizer to the crop. Some growers prefer black woven landscape fabric, which can be reused for seven or more years. Weeds emerging through planting holes are removed manually, and alley weeds are managed by hoeing, cultivation, mowing, organic mulch, or cover cropping.

NOTE: When plastic or other synthetic mulches are used for organic production, they must be removed from the field at the end of the growing or harvest season.

Tomatoes in black plastic
Figure 2. (a) Vegetable growers setting tomato starts into black plastic mulch. (b) Tomato at flowering in black plastic, with hay mulch in alleys. Photo credits: (a) Becky Crouse, Marketing Manager, Potomac Vegetable Farms, Purcellville, VA; (b) Mark Schonbeck, Virginia Association for Biological Farming.

Because tomato yield and quality can suffer from excessive soil heating, some growers spread straw or hay over black plastic mulch when summer heat arrives, or use a light-colored plastic for later plantings. In hot climates, a light-reflecting film mulch may be appropriate for main-season or late plantings of pepper and eggplant as well (Fig. 3).

Light-reflecting film mulch
Figure 3. This light-reflecting film mulch (opaque polyethylene with an aluminized surface), results in lower soil temperatures than black plastic, and repels aphids and some other pests from the pepper crop. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

If plastic mulch is not used, farmers either hoe or cultivate the crop as needed through the minimum weed-free period, or cultivate once or twice, then spread an organic mulch, such as straw or hay, to delay weed emergence until the crop is fully established (Fig. 4). Because organic mulch conserves moisture and moderates soil temperature, it can enhance tomato yields if applied after the soil has warmed sufficiently (Schonbeck and Evanylo, 1998).

Straw mulch for tomatoes, peppers, and eggplant.
Figure 4. (a) Tomato in hay mulch passed through its minimum weed-free period before weeds began to break through the mulch. (b) Straw mulch, applied after cultivation and when soil temperatures were near optimal for eggplant and pepper, has given excellent weed control. Crops were planted in double rows, and the eggplant has closed canopy, thus shading out late-emerging weeds. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Tall tomato varieties grown on trellises or tall stakes can cast sufficient shade to hinder between-row weed growth. However, in moist conditions, the tomato crop may lose its foliage to diseases toward the end of the harvest period, which allows weed growth to resume.

Most Serious Weeds in Solanaceous Crops in the Southern Region

Some of the most widespread and troublesome weeds in solanaceous crops in the Southern region include yellow nutsedge (Cyperus esculentus), purple nutsedge (C. rotundus), morning glories (Ipomoea spp.), and pigweeds (Amaranthus spp.) (Fig. 5) (Webster, 2006). Bermuda grass (Cynodon dactylon), johnsongrass (Sorghum halapense), crabgrasses (Digitaria spp.), foxtails (Setaria spp.), and galinsoga (Galinsoga spp.) have also been cited as major weeds of tomato family in some southern states.

Troublesome weeds for solanaceous crops
Figure 5. Some troublesome weeds for solanaceous crops: (a) Yellow nutsedge and two species of morning glory emerging in alley between tomato rows. (b) Purple nutsedge competing severely against pepper. (c) late-season morning glories climbing mature tomato plants, with pigweeds growing in alleys (foreground). Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Morning glories are especially troublesome because they can emerge through six inches of straw mulch, readily grow toward and emerge through planting holes in plastic mulches, and climb crop plants. The fast-growing, entangling vines interfere with harvest and can smother the crop.

The sharp-pointed shoots of emerging nutsedges and Bermuda grass can penetrate black plastic film or landscape fabric. The weeds then compete with the crop, and complicate end-of-season mulch removal.

Pigweeds compete aggressively against solanaceous crops, owing to their tall stature and rapid growth during hot weather. Later-emerging pigweed can cause problems, even when the crop is large enough to shade out most grassy weeds and nutsedges.

Horsenettle (Solanum carolinense) (Fig. 6a), black nightshade (Solanum nigrum) and other solanaceous weeds are alternate hosts for diseases such as early blight (Alternaria solani), septoria leaf spot (Septoria lycopersici), and late blight (Phytophthora infestans) of tomato, and phytophthora blight (P. capsici) of pepper. Many weed species, including winter annuals like common chickweed (Stellaria media) (Fig. 6b), mouse ear chickweed (Cerastium vulgatum), cutleaf evening primrose (Oenothera laciniata), and cudweeds (Gnaphalium spp.) can harbor tomato spotted wilt virus (TSWV) and thrips (Thysanoptera), an insect pest that acts as a vector (carrier) and transmits the virus from weed to crop (Louws, 2009; Martinez, 2008). TSWV is most serious in areas with mild winters that do not kill off thrips populations.

Horsenettle and common chickweed
Figure 6. Two weeds that pose a disease and pest hazard to solanaceous crops. (a) Horsenettle hosts early blight and septoria leaf spot of tomato, phytophthora blight of pepper, and false potato beetle, which can severely damage eggplant. (b) Common chickweed can harbor the tomato spotted wilt virus (TSWV) and its thrips vector over the winter, thereby propagating the disease from one season to the next. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Preventive Weed Management in Solanaceous Crops

Preventive weed management begins with planning and field preparation. If possible, avoid planting tomato, pepper, and eggplant in fields with high populations of nutsedge, Bermuda grass, or other aggressive weeds. Use stale seedbed prior to planting to reduce seed populations of pigweed, morning glory, and other summer annual weeds. Weeds known to carry a disease or pest of solanaceous crops that is prevalent in the area should be well controlled before rotating the field to tomato or other crops in this plant family.

A good crop rotation that includes weed-smothering cover crops can reduce weed problems in tomato, pepper, and eggplant (Diver et al., 2007). Alternate warm- and cool-season vegetables in successive years, and grow competitive summer cover crops like buckwheat, cowpea, and sorghum–sudangrass during the season prior to tomato family production. Rotate heavily weed-infested fields into a perennial grass–clover sod for 1–3 years to reduce the weed seed bank.

Optimize soil temperature for crop establishment. For early plantings, use black plastic or fabric mulches, low or high tunnels (Fig. 7), or other strategies to raise soil temperature, so that crop development is not delayed by cold soil. Be sure the soil is thoroughly warm, (~70 °F) before transplanting eggplant or hot pepper. Delay application of straw or other soil-cooling mulches near plants until the soil has reached optimal temperatures. For late-season tomato plantings during hot weather, use a white or reflective film mulch (for best weed control, use an opaque white-on-black film, or coat black plastic with a nontoxic whitewash), or apply straw soon after planting to limit solar heating of the soil.

Companion-cropped high tunnel tomatoes
Figure 7. Tomato planted in March in a high tunnel will begin producing fruit by June at Dayspring Farm in the Tidewater region of Virginia. The companion crops of lettuce and bok choi have limited early-season weed growth, and will soon be harvested. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Use slow-release organic nutrient sources to meet crop nutritional needs. Avoid more soluble, faster-releasing fertilizers, which could stimulate early-season growth of pigweeds, black nightshade, and other N-responsive weeds. On good, biologically active soil, tomato N needs can be fully met with legume cover crops and compost, and yield better with these N sources than with soluble fertilizers (Diver et al., 2007).

Use in-row drip irrigation to deliver water and liquid organic fertilizers (if needed) preferentially to the crop (Fig. 8). Subsurface drip irrigation (lines buried several inches deep in the crop row) provides moisture to crop roots while leaving the soil surface dry and thereby deterring weed seed germination.

In-row drop irrigation of tomatoes
Figure 8. In-row drip irrigation can give the crop the edge over weeds, especially in a dry season. A mulch application would further enhance the crop's advantage. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Transplant the crop immediately after preparing the bed, especially if plastic mulch is not laid before planting. Even a couple days' delay can give the weeds a significant head start. For tomato, choose row spacing, plant spacing, and method of support that optimizes air circulation for disease control, and facilitates cultivation or mulching. For pepper and eggplant, where disease pressure is light, plant a staggered double row at a plant spacing that promotes canopy closure within the bed or grow zone without unduly crowding the plants.

Cultivation and Other Weed Control Tactics

If plastic film or fabric mulch is not used, cultivate or hoe around the plants when the first flush of weeds is less than one inch tall. Cultivate shallowly to avoid root pruning, especially in pepper. If crop plants are large enough and the southern stem blight pathogen is not present in the soil, adjust cultivation implements to throw soil into the crop rows to bury small within-row weeds, rather than trying to sever or uproot them.

Without mulch, as many as three cultivations may be needed before the end of the minimum weed-free period. Many organic growers hoe or cultivate once or twice to remove early weeds, then apply 3–4 inches of straw, hay, or other organic mulch. This approach conserves soil moisture, adds organic matter, prevents soil splash during rains, and can provide excellent weed control in fields that are not heavily infested with aggressive perennial weeds or morning glories.

For tomato, pepper, and eggplant transplanted into plastic film or fabric mulch, some manual labor is usually needed to remove weeds that emerge through planting holes. Usually, one manual weeding is sufficient, after which the growing crop shades out emerging weeds. Control alley weeds by cultivation, hoeing, mowing, or spreading straw or other organic mulch in alleys and overlapping edges of the plastic.

Once the crop has passed through its minimum weed-free period, manage later-season weeds so that they do not hinder air circulation and promote foliar diseases (Fig. 9), interfere with harvest, or propagate themselves. Pull or cut morning glories and other vining weeds before they begin to climb the crop. Remove large "escapes" before they set seed. Hoe, cultivate or mow closely any nutsedge or other invasive perennials to disrupt formation of new rhizomes and tubers.

Late-season weeds in tomatoes
Figure 9. These weeds emerged late enough not to compete directly with the established tomato crop; however, they reduced air circulation and promoted the development of fungal diseases, which have defoliated the lower parts of these plants. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Additional Weed Management Strategies

Winter cover crops, such as cereal rye–hairy vetch or barley–crimson clover, can be roll-crimped or flail-mowed for no-till planting of summer vegetables. The rye–vetch mulch appears especially promising for tomato production. In a study in Beltsville, MD, tomato planted in mow-killed hairy vetch yielded about 20% more than tomato in black plastic (Abdul-Baki and Teasdale, 1993, 1997). Mowed rye mulch can suppress weeds for 4–8 weeks without adversely affecting tomato yield (Smeda and Weller, 1996) and vetch residues have been shown to enhance disease resistance, prolong active photosynthesis and delay leaf senescence in tomato by modifying tomato gene expression (Kumar et al., 2004). No-till planting into mechanically killed cover crop is recommended for main-season and late tomato plantings, but not early plantings, as the cover crop mulch may delay soil warming, crop establishment, and fruit ripening.

Winter cover crops can be strip-tilled several weeks before tomato or pepper planting to promote soil warming in the crop grow-zone. Cover crops growing in alleys can be maintained by mowing to keep alley weeds down (Fig. 10); in some cases, the mowings can provide clean mulch for the vegetable. Season-long living mulches of white clover, subterranean clover, or ryegrass, managed by mowing or partial tillage, have also been used successfully for alley weed control in tomato (Diver et al., 2007). However, clovers host nematode pests that affect tomato, such as root-knot nematode; thus, clovers should not be grown with or immediately before solanaceous vegetables in fields in which these nematode pests are present.

Strip-tilled rye before transplanting tomatoes
Figure 10. A winter rye cover crop was strip-tilled prior to transplanting tomato at Seven Springs Farm in Check, VA (Appalachian region). The cover crop, maintained by mowing, has suppressed alley weeds. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Finally, cover crops can be planted in alleys between beds several weeks after vegetable planting (relay intercropping) to suppress weeds, protect soil, and add organic matter. Examples include buckwheat, cowpea, millet, or white clover, mowed if needed to maintain air circulation around the crop.

References Cited
  • Abdul-Baki, A. A., and J. R. Teasdale. 1993. A no-tillage tomato production system using hairy vetch and subterranean clover mulches. HortScience 28: 106–108.
  • Abdul-Baki, A. A., and J. Teasdale. 1997. Sustainable production of fresh market tomatoes and other summer vegetables with organic mulches. Farmers' Bulletin No. 2279. USDA–Agriculture Research Service, Washington, D.C. (Available online at: http://www.ars.usda.gov/is/np/SustainableTomatoes2007/SustainableTomatoes2007Intro.htm) (verified 10 Sept 2012).
  • Diver, S., G. Kuepper, and H. Born. 2007. Organic tomato production [Online]. Available at: https://attra.ncat.org/attra-pub/summaries/summary.php?pub=33 (verified 10 Sept 2012).
  • Kumar, V., D. J. Mills, J. D. Anderson, and A. K. Mattoo. 2004. An alternative agriculture system is defined by a distinct expression profile of select gene transcripts and proteins. Proceedings of the National Academy of Sciences 101: 10535–10540. (Available online at: http://dx.doi.org/10.1073/pnas.0403496101) (verified 10 Sept 2012).
  • Louws, F. 2009. Managing vegetable diseases. Presentation at the 24th annual Sustainable Agriculture Conference of the Carolina Farm Stewardship Association, Black Mountain, NC, Dec 5, 2009.
  • Martinez, N. 2008. Tospoviruses in Solanaceae and other crops in the coastal plain of Georgia: Epidemiology, weed hosts. University of Georgia. (Available online at: http://www.caes.uga.edu/topics/diseases/tswv/vegcrops/tospoviruses/epidemiologywh.html) (verified 10 Sept 2012).
  • Monks, D. 1993. Veg-I-News. Cooperative Extension Service, North Carolina State University. Vol. 12, No. 4.
  • Peet, M. 1996. Sustainable practices for vegetable production in the South. Focus Publishing, R. Pullins Company, Newburyport, MA.
  • Riggs, D.I.M., R. R. Bellinder, and R. W. Wallace. 1991. The effect of one, two and three month weed-free periods on yield of late season tomatoes. HortScience 26: 152. (Available online at: http://hortsci.ashspublications.org/content/26/6/768.4.abstract) (verified 10 Sept 2012).
  • Schonbeck, M. W., and G. E. Evalylo. 1998. Effects of mulches on soil properties and tomato production. I. Soil temperature, soil moisture, and marketable yield. Journal of Sustainable Agriculture 13: 55–81. (Available online at: http://dx.doi.org/10.1300/J064v13n01_06) (verified 10 Sept 2012).
  • Smeda, R. J., and S. C. Weller. 1996. Potential of rye (Secale cereale) for weed management in transplant tomatoes (Lycopersicon esculentum). Weed Science 44: 596–602. (Available online at: http://www.jstor.org/stable/4045642) (verified 10 Sept 2012).
  • Swaider, J. M, G. W. Ware, and J. P. McCollum. 1992. Producing vegetable crops, 4th ed. Interstate Publishers, Inc, Danville, IL.
  • Teasdale, J. R., and D. Colaccicco. 1985. Weed control systems for fresh market tomato production on small farms. Journal of the American Society of Horticultural Science 110: 533–537.
  • Tindall, J. A., H. A. Mills, and D. E. Radcliffe. 1990. The effect of root zone temperature on nutrient uptake of tomato. Journal of Plant Nutrition 13: 939–956. (Available online at: http://dx.doi.org/10.1080/01904169009364127) (verified 10 Sept 2012).
  • Weaver, S. E., and C. S. Tan. 1983. Critical period of weed interference in transplanted tomatoes (Lycopersicon esculentum): Growth analysis. Weed Science 31: 476–481. (Available online at: http://www.jstor.org/stable/4043595) (verified 10 Sept 2012).
  • Webster, T. M. 2006. Weed survey – southern states. Vegetable, fruit and nut crops subsection. Proceedings of the Southern Weed Science Society 59: 260–277. (Available online at: http://www.swss.ws/NewWebDesign/Publications/Weed%20Survey%20Archives/Southern%20Weed%20Survey%202006%20Vegetables%20and%20Fruits.pdf) (verified 10 Sept 2012).

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 4873

Current and Past Issues of eOrganic Updates Newsletter

lun, 2012/09/10 - 18:08

Subscribe to the eOrganic Updates Newsletter

Issues

September 2012

August 2012

June 2012

May 2012

Annual report 2011

December 2011

September 2011

August 2011

July 2011

May 2011

September 2010

August 2010

July 2010

June 2010

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 6127

September 2012 eOrganic Newsletter

lun, 2012/09/10 - 18:01
In this Issue
  • What's New at eOrganic
  • New Video: What is Organic?
  • New Farmer's Guide to Organic Contracts
  • New Organic Literacy Initiative by the NOP
  • NOP News: Last Call for Certification Cost Share
  • NCAT Webinar on Organic Conservation Practice Standards
  • CCOF Upcoming Webinars and Events
What's New at eOrganic

Organic Mulching Materials for Weed Management, by Mark Schonbeck, Virginia Association for Biological Farming. Available at http://www.extension.org/pages/65025

Synthetic Mulching Materials for Weed Management, by Mark Schonbeck, Virginia Association for Biological Farming. Availble at http://www.extension.org/pages/65191

Video: Healthy Cow Check-Up—How to Perform a Physical Exam. Dr. Hubert Karreman, Penn Dutch Cow Care. Available at http://www.extension.org/pages/64752

Organic News and Announcements New Video: What is Organic?

A new video filmed for the St. Paul, farmer's market features eOrganic certification group leader Jim Riddle, who addresses the question "What is Organic" from his Minnesota farm.  Watch the video at http://www.youtube.com/watch?v=BQskVbWGT2c

New Farmer's Guide to Organic Contracts Available Online

The goal of FLAG's new Farmers' Guide to Organic Contracts is to help organic farmers make informed decisions when evaluating, negotiating, and managing contract agreements with buyers of organic farm products. This farmer-friendly guide includes: 1) a basic overview of contract laws important to farmers; 2) a Quick Organic Contract checklist and practical toolkit farmers can use to review contract offers; 3) highlighted sections showing how NOP regulations interact with organic contracts; 4) explanations and examples of over 100 types of organic contract provisions; and 5) detailed information about solving the types of contract disputes that commonly arise in the organic market. The guide is available for free download at http://www.flaginc.org.

New Organic Literacy Initiative by the NOP

On September 4, the U.S. Department of Agriculture released a series of resources as part of its new Organic Literacy Initiative, an effort to help connect current and prospective organic farmers, ranchers, and processors with relevant USDA resources. Find all the publications at the new Organic Literacy Home Page. The purpose of the Organic Literacy Initiative is to provide USDA staff as well as organic producers and handlers with detailed and consistent information about organic agriculture and the programs and services USDA offers to support it. One of the goals of the initiative is to help USDA staff around the U.S. be better equipped to help farmers, ranchers, and processors understand organic certification and access relevant USDA services.

The materials available include:

An Organic 101 training about what the organic label means and how certification works;
A brochure that contains information about organic standards and certification and a brief description of USDA resources;
An Organic Resource Guide that outlines how each USDA agency supports organic agriculture and provides relevant USDA contact information; and
A USDA blog that highlights organic topics.

NOP News: Last call for Organic Certification Cost Share

Funds are still available for the 2012 Organic Certification Cost Share Program—as much as $750 per certified operation—to certified organic farmers and businesses to help cover the cost of organic certification. Newly certified applicants must have an organic certificate dated between October 1, 2011 and September 30, 2012 to apply during the 2012 funding cycle. If you are renewing your certification, you can submit your application as soon as you pay your certification fees. Contact your state’s department of agriculture for an application. You can find names and phone numbers at http://www.ams.usda.gov/NOPCostShareProgramParticipants, and some states have forms online for you to download. Your state’s deadline may be as early as September 30th, so don’t wait—apply today!

NCAT Webinar on Conservation Practice Standards on September 27, 2012

NCAT is hosting a webinar on September 27, 2012 at 1-2 PM Eastern Time on the Links between Biodiversity Requirements of Organic Systems and NRCS Practice Standards.  The webinar is funded by an NRCS Conservation Innovation Grant. Register at https://www2.gotomeeting.com/register/359336938.

Biodiversity conservation is part of the definition of organic farming, and the NOP requires that farmers and ranchers maintain or improve their soil, water, wetlands, woodlands, and wildlife. In addition, seven other NOP regulations relate to biodiversity and natural resource conservation. NRCS Conservation Practice Standards that help operators meet these NOP requirements will be discussed, including those protecting resources, providing conservation buffers, and supporting wildlife habitat. Also presented will be examples of practices used by organic farmers to maintain or enhance natural resources on their operations. Presenters: Jo Ann Baumgartner, Wild Farm Alliance, Jim Riddle, University of Minnesota, and Tom Broz, Live Earth Farms.

CCOF Upcoming Webinars and Events

California Certified Organic Farmers is hosting several marketing webinars this fall: Marketing 101 on September 26, and Sales Basics on October 3rd—as well as an in-person, all-day Organic Wholesale Market Tour on October 16 in San Francisco! Register early since space is limited. Find out more information on their Education and Events page at http://ccof.org/programs.php

eOrganic Mission

eOrganic is a web community where organic agriculture farmers, researchers, and educators network; exchange objective, research- and experience-based information; learn together; and communicate regionally, nationally, and internationally.

eOrganic Resources

Find all eOrganic articles, videos and webinars at http://extension.org/organic_production

Connect with eOrganic on Facebook and Twitter, and subscribe to our YouTube channel!

Have a question about organic farming? Use the eXtension Ask an Expert tool to connect with eOrganic. Tag your question as "organic production" to make sure it reaches our members.

Spread the word! If you would like eOrganic literature to hand out at your next conference or workshop, please get in touch, and use our online form to request materials.

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 8211

eOrganic Updates

lun, 2012/09/10 - 17:44

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T1123

Can I Use This Input on My Organic Farm?

mar, 2012/09/04 - 13:43
imageUsing an input that is not approved can cost you your organic certification. Learn how to determine if a synthetic or natural material can be used in organic production.

eOrganic authors:

Nick Andrews, Oregon State University

Brian Baker, Organic Materials Research Institute

Jim Riddle, University of Minnesota

Assessing Inputs

The National Organic Program (NOP) final rule (United States Department of Agriculture [USDA], 2000) emphasizes the use of preventive and cultural methods such as crop rotation, cover cropping, sanitation measures, and nutritious feed rations to build soil fertility, prevent pest problems, and maintain livestock health. In fact, the NOP requires that management practices to prevent pests, weeds and diseases, including soil-building crop rotations; sanitation measures to remove disease vectors, weed seeds, and pest habitat; selection of site-suitable and resistant plant and livestock species and varieties; release of pest predators and parasites; development of habitat for pest predators; lures, traps and repellants; mulching, mowing, grazing, mechanical, flame, and/or hand weeding; and cultural practices to prevent weed, pest and disease problems must be implemented, and found to be insufficient, prior to the use of any input.

Many organic farmers save money and produce high-quality crops with little or no off-farm inputs, but most producers rely on at least some purchased inputs. Purchasing inputs brings up the question: Is this product allowed? Given that most agricultural inputs are not produced with NOP standards in mind, those of us trying to meet the standards have to ensure that we use only approved products.

With full implementation of the National Organic Program regulations on October 21, 2002, it’s the National Organic Program (NOP) that allows or prohibits a material, via the National List of Allowed and Prohibited Substances (National List). This is a generic materials list. The NOP does not publish a brand name materials list. Brand name lists evaluate a formulated product’s compliance to the requirements of the National List, but do not carry any regulatory weight. Of course, you are free to use brand name lists for guidance (as most organic certifiers do), but certification agencies are responsible for evaluating materials to be used by producers and handlers for compliance with the National List requirements. As a producer, this means that you must submit a list of all inputs you use or intend to use as part of your Organic System Plan, and your certification agency will determine if the substances are allowed for organic production.

Making Sense of the National List

The portion of the NOP concerned with the National List begins at section 205.105 (see “NOP on materials” below). Simply put, it says we cannot use products with synthetic ingredients for orgsanic crop or livestock production, unless they are specifically allowed and appear on the National List (see definition in “NOP on materials”). Some nonsynthetic (natural) substances are also prohibited (see sections 205.602 and 205.604). In other words, synthetic materials cannot be used unless they are specifically approved, and natural materials can be used unless they are specifically prohibited. The National List specifies the allowed synthetic substances and prohibited nonsynthetic substances (see section 205.601), along with specific restrictions, or annotations, regarding the source or use of the substance. The National List doesn’t include numerous natural, nonsynthetic substances, such as gypsum, limestone, or rock phosphate, which are allowed by definition.

Reading Labels

Next stop is the product label. If the ingredients are nonsynthetic and not included as prohibited in section 205.602, they are allowed. If there are synthetic ingredients, check to see if they are specifically allowed. Be sure to check section 205.601 for crop and 205.603 for livestock annotations. These annotations are where I’ve seen some challenges for growers. The annotations often regulate or place certain restrictions on the manufacturing processes or the use of a material. For example, Lidocaine is allowed as a local anesthetic, but its use requires a withdrawal period of 90 days for livestock intended for slaughter and seven days for dairy animals. Compliance with restrictions must be documented when an annotated material is used.

Reading the label may not be enough to determine if a product complies with National List annotations. Labels frequently do not provide all the information about the manufacturing process. For example, liquid fish products are allowed as plant or soil amendments (see §205.601.j.7). They “can be pH adjusted with sulfuric, citric or phosphoric acid. The amount of acid used shall not exceed the minimum needed to lower the pH to 3.5.” Ingredient lists on liquid fish do not address this issue. Before purchasing or using a liquid fish product, a grower should contact the manufacturer or confirm that the product is listed on a brand name list of NOP compliant products.

Non-active or inert ingredients in pesticide formulations are classified according to the level of toxicological concern. EPA has changed how it lists inert ingredients, and the NOP has taken over the maintenance of the list of substances used as inert ingredients that EPA determined to be of minimal concern prior to 2004. To be NOP compliant, all synthetic inert ingredients in pesticides must be classified as minimum risk, appear specifically on the National List, or be used in passive pheromone dispensers. Inert ingredients do not appear on labels, so verifying compliance with this annotation requires the cooperation of the pesticide registrant.

If you contact manufacturers, try to get answers in writing or at least record what you learn. If you are unsure of the information that you need, contact your certifier for guidance and they should be able to help you ask the right questions. When considering a new product, be sure to plan ahead. In trickier cases, a simple “yes” or “no” answer may not be possible over the phone.

As the saying goes, “The devil is in the details.” It’s these tricky cases that cause the headaches. It’s tempting to just refer to a brand name materials list and use those products. But remember, just because a product is not on a particular brand name materials list does not necessarily mean that it is prohibited by the NOP. Also remember that only the NOP carries regulatory weight; all other lists are based on evaluations of compliance to the rule. 

Brand Name Material Lists

The NOP established a policy that each Accredited Certifying Agent (ACA, certifier) is responsible for conducting its own reviews of inputs for agricultural production, such as formulated pesticides and soil amendments. The NOP also allows certifying agents to recognize reviews conducted by other certifying agents and competent third-party reviewers as described in a letter to organic certifiers on verification of materials (Robinson and Bradley, 2008), and later confirmed by a Policy Memo in the NOP Policy Handbook. All ACAs are required to verify, along with their clients, that all materials used or planned for use by certified organic operations comply with the NOP. To paraphrase, ACAs have three options available to determine whether branded or formulated products comply:

  1. ACAs can contact the manufacturer to obtain disclosure of the contents of the product and verify that they all comply;
  2. ACAs may consult with another ACA that has reviewed the information and accept their determination that the material is NOP compliant; or
  3. ACAs may consult with a reputable third party source, such as the Environmental Protection Agency (EPA) or the Organic Materials Review Institute (OMRI), that reviews materials for compliance with the NOP regulation.

ACAs must document their determinations and verify that the inputs are used according to the regulation. ACAs must either have the capacity and expertise to review products, or contract with organizations accredited do so. Many ACAs contract with OMRI, a non-profit initially established by certifiers specifically for that purpose. The Washington State Department of Agriculture (WSDA) also reviews products according to the NOP and publishes a list of brand name products that other ACAs use. These lists are not comprehensive, so there may be other brand name products that can be used. However, in order to be sure that a product complies, the manufacturer must fully disclose all ingredients and manufacturing processes to an ACA or a third party contracted by the ACA. All ingredients must comply with the standards described above.

One Step at a Time

Before using a new product, check for recent OMRI or WSDA approval of the product. If it isn’t listed, follow these steps:

  1. Evaluate each label ingredient for compliance with the NOP and any annotations on the National List. The OMRI Generic Materials List may also be helpful.
  2. Contact the manufacturer if necessary.
  3. Document compliance with all NOP crop and livestock annotations.

Since this process can take some time, be sure to plan ahead when developing or updating your Organic System Plan (OSP). Keep records of all communications with input manufacturers, certifiers, and input review services. Keep labels, receipts, shipping invoices, and input application records. The documentation required to demonstrate compliance with the NOP can seem daunting and sometimes takes time away from working the land. However, this careful verification gives organic consumers confidence in the organic standard they have grown to trust.

NOP Citations on Materials

§ 205.105 Allowed and prohibited substances, methods, and ingredients in organic production and handling.
To be sold or labeled as “100 percent organic,” “organic,” or “made with organic (specified ingredients or food group(s)),” the product must be produced and handled without the use of:
(a) Synthetic substances and ingredients, except as provided in § 205.601 or § 205.603;
(b) Nonsynthetic substances prohibited in § 205.602 or § 205.604;
(c) Nonagricultural substances used in or on processed products, except as otherwise provided in § 205.605;
(d) Nonorganic agricultural substances used in or on processed products, except as otherwise provided in § 205.606;
(e) Excluded methods, except for vaccines, provided that the vaccines are approved in accordance with § 205.600(a);
(f) Ionizing radiation, as described in Food and Drug Administration regulation, 21 CFR 179.26; and
(g) Sewage sludge.

§ 205.206 Crop pest, weed, and disease management practice standard
(e) When the practices provided for in paragraphs (a) through (d) of this section are insufficient to prevent or control crop pests, weeds, and diseases, a biological or botantical substance or a substance included on the National List of synthetic substances allowed for use in organic crop production may be applied to prevent, suppress, or control pests, weeds, or diseases; Provided, That, the conditions for using the substance are documented in the organic system plan.

§ 205.602 Synthetic substances allowed for use in organic crop production.
In accordance with restrictions specified in this section, the following synthetic substances may be used in organic crop production: Provided, That, use of such substances do not contribute to contamination of crops, soil, or water. Substances allowed by this section, except disinfectants and sanitizers in paragraph (a) and those substances in paragraphs (c), (j), (k), and (l) of this section, may only be used when the provisions set forth in §205.206(a) through (d) prove insufficient to prevent or control the target pest.

  § 205.602 Nonsynthetic substances prohibited for use in organic crop production.
The following nonsynthetic substances may not be used in organic crop production:
(a) Ash from manure burning
(b) Arsenic
(c) Lead salts
(d) Sodium fluoaluminate (mined)
(e) Strychnine
(f) Tobacco dust (nicotine sulfate)
(g) Potassium chloride—unless derived from a mined source and applied in a manner that minimizes chloride accumulation in the soil.
(h) Sodium nitrate—unless use is restricted to no more than 20 percent of the crop’s total nitrogen requirement.
(i-z) [Reserved]

§ 205.604 Nonsynthetic substances prohibited for use in organic livestock production.
The following nonsynthetic substances may not be used in organic livestock production:
(a) Strychnine
(b)-(z) [Reserved]

NOP definition of "synthetic" -  "A substance that is formulated or manufactured by a chemical process that chemically changes a substance extracted from naturally occurring plant, animal or mineral sources, except that such term shall not apply to substances created by naturally occurring biological processes."

References and Citations Additional Resources

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 2473

Organic Vegetable Production Systems, Soil and Fertility Management in Organic Farming Systems

mar, 2012/09/04 - 12:24

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T879,867

Soil and Fertility Management in Organic Farming Systems

mar, 2012/09/04 - 12:24

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T867

Soil Nematodes in Organic Farming Systems

mar, 2012/09/04 - 12:21

eOrganic authors:

Carmen Ugarte, University of Illinois

Ed Zaborski, University of Illinois

Introduction

Nematodes are microscopic, wormlike organisms (Fig. 1) that live in water films and water-filled pore spaces in the soil. Typically, they are most abundant in the upper soil layers where organic matter, plant roots, and other resources are most abundant. Nematode abundance in soils—managed and unmanaged—ranges from 1–10 million individuals/m2 (Peterson and Luxton, 1982; Lavelle and Spain, 2001).

Figure 1. A typical free-living, bacterial-feeding nematode.
Figure 1. A typical free-living, bacterial-feeding nematode, less than 1mm (0.04in) in length. Figure credit: Ed Zaborski, University of Illinois.

Most research on soil nematodes has focused on the plant-parasitic nematodes that attack the roots of cultivated crops. Less attention has been given to nematodes that are not plant-feeders and play beneficial roles in the soil environment. This article describes the important roles played by nematodes in soil ecosystems, as well as their potential to be used as indicators of soil condition in organic farming systems.

Nematode Feeding Habits

Nematodes can be classified into functional groups based on their feeding habits, which can often be deduced from the structure of their mouthparts (Fig. 2). In agricultural soils, the most common groups of nematodes are the bacterial-feeders, fungal-feeders, plant parasites, predators, and omnivores. Predatory nematodes feed on protozoa and other soil nematodes. Omnivores feed on different foods depending on environmental conditions and food availability; for example, omnivorous nematodes can be predators, but in the absence of their primary food source, they can feed on fungi or bacteria.

Figure 2. Nematode feeding types; mouthpart structures.
Figure 2. Nematodes can be classified into different feeding groups based on the structure of their mouthparts. (a) bacterial feeder, (b) fungal feeder, (c) plant feeder, (d) predator, (e) omnivore. Figure credit: Ed Zaborski, University of Illinois.

Importance of Nematodes in Agricultural Systems

Nematodes contribute to a variety of functions within the soil system. In agricultural systems, nematodes can enhance nutrient mineralization and act as biological control agents.

Nematodes and Soil Fertility

Soil nematodes, especially bacterial- and fungal-feeding nematodes, can contribute to maintaining adequate levels of plant-available N in farming systems relying on organic sources of fertility (Ferris et al., 1998). The process of converting nutrients from organic to inorganic form is termed mineralization; mineralization is a critical soil process because plants take up nutrients from the soil primarily in inorganic forms. Nematodes contribute directly to nutrient mineralization through their feeding interactions. For example, bacterial-feeding nematodes consume N in the form of proteins and other N-containing compounds in bacterial tissues and release excess N in the form of ammonium, which is readily available for plant use. Indirectly, nematodes enhance decomposition and nutrient cycling by grazing and rejuvenating old, inactive bacterial and fungal colonies, and by spreading bacteria and fungi to newly available organic residues. In the absence of grazers, such as nematodes and protozoa, nutrients can remain immobilized and unavailable for plant uptake in bacterial and fungal biomass.

Bacterial-feeding nematodes are the most abundant nematode group in agricultural soils. Their abundance closely follows that of bacterial populations, which tend to increase when soil disturbances, such as tillage, increase the availability of readily-decomposable organic matter. Nitrogen mineralization in the soil occurs at a higher rate when bacterial-feeding nematodes are present than when they are absent. The contribution of bacterial-feeding nematodes to soil N supply depends, in part, on the quality and quantity of soil organic matter fueling the system. Net N mineralization from decomposing organic residues takes place when the carbon:nitrogen (C:N) ratio of organic residue is below 20 (that is, 20 parts C to 1 part N). When the C:N ratio is greater than 30, the rate of mineralization decreases because microbes compete for N to meet their nutritional requirements. In this situation, N is immobilized in the microbial biomass. Incorporation of manure, compost, and cover crops with intermediate C:N ratios (ranging from 10 to 18) may stimulate bacterial growth and the abundance of bacterial-feeding nematodes, and increase soil N availability to plants.

Fungal-feeding nematodes are relatively more abundant in less-disturbed (e.g. notill systems) and perennial systems, where conditions for fungal growth are promoted, than in disturbed systems. Like bacterial feeding nematodes, fungal-feeding nematodes contribute to the process of nutrient mineralization by releasing N and other plant nutrients from consumed fungal tissue. However, in agricultural systems, bacterial-feeding nematodes typically release more inorganic N than fungal-feeding nematodes.

Nematodes as Natural Enemies and Biological Control Agents

Predatory nematodes are of interest because of their role in regulating the populations of other organisms. They generally feed on smaller organisms like protozoa and other nematodes. Thus they can help moderate population growth of bacterial- and fungal-feeding nematodes and protozoa, and help regulate populations of plant-parasitic nematodes.

Insect-parasitic nematodes are species of bacterial-feeding nematodes that live in close association with specific species of bacteria; together, they can infect and kill a range of insect hosts. The infective juvenile stage of insect-parasitic nematodes seeks out insect hosts to continue its development into adults. Once a host is found, the nematodes penetrate the insect body and release their bacterial associates into the insect’s body cavity. These bacteria multiply and overwhelm the immune response of the host insect, ultimately killing the host. The nematodes feed on these bacteria, mature, and reproduce until all the resources within the insect host are consumed; then, infective juvenile nematodes escape the insect host's body and disperse in the soil to seek new hosts. Insect-parasitic nematodes are available commercially for use in inundative releases to manage the populations of a variety of insect pests.

Plant-Parasitic Nematodes

Most plant-parasitic nematodes feed on the roots of plants. Some species attach to the outside surface of plant roots (Fig. 3), piercing the root tissue to suck up the cellular content; other species pierce and penetrate the roots of plants, living and reproducing entirely within the root itself. A relatively small number of important plant-parasitic nematode species are known to cause substantial economic damage in cropping systems around the world. The determination of tolerance limits or economic thresholds for plant-parasitic nematodes varies with many factors like species, plant tolerance, and soil type. Because plant parasitic nematodes show varying degrees of host specificity, carefully designed crop rotations are usually a powerful tool for reducing nematode-associated yield losses.

Figure 3. Symptoms of white potato cyst nematode, Globodera pallida.
Figure 3. White potato cyst nematode, Globodera pallida (Stone) Behrens, on plant roots. Cyst nematode females attach to root systems with their mouthparts to feed, and then their bodies swell into egg-filled cysts that can be visible to the naked eye. Figure credit: Bonsak Hammeraas, Bioforsk—Norwegian Institute for Agricultural and Environmental Research, Bugwood.org.

Soil Nematode Communities

The proportions of the different feeding groups in the soil nematode community vary between systems and seasons, and they are influenced by a variety of factors, including crop and soil management practices (Freckman and Ettema, 1993) and the presence and abundance of natural enemies. Management practices like tillage, crop rotation, and the use of organic amendments influence the physical and biological characteristics of the soil that influence the abundance of nematodes. Fungal-feeding, predatory, and omnivorous nematodes are very sensitive to soil disturbances (Ferris et al., 2001), and agricultural systems with fewer physical and chemical disturbances, such as pastures, hay fields, and orchards, tend to support larger populations of these nematodes than more frequently disturbed systems like vegetable- and row-crop fields. On the other hand, tillage and incorporation of organic residues increase the proportion of some bacterial-feeding nematodes (Griffiths et al., 1994; Ferris et al., 2001; Nahar et al., 2006), often offsetting declines in the numbers of other feeding groups and increasing the total abundance of nematodes (Neher, 1999). The wide variety of natural enemies that feed on or infect nematodes—predatory nematodes, predatory microarthropods, and nematode-trapping fungi, for example—may have a considerable impact on nematodes in agricultural systems (Stirling, 1991).

Implications for Farming System Management

Agricultural management may increase the abundance of soil nematodes, primarily through the increase in the abundance of bacterial-feeding nematodes associated with tillage and the incorporation of organic residues (Neher, 1999). Soil conditions in agricultural production systems can be improved by enhancing nutrient availability and providing habitat for beneficial soil organisms. Maintenance of large populations of bacterial-feeding nematodes with practices that promote N mineralization throughout the growing season may enhance crop productivity, but a surplus of mineral N is not desirable from the environmental point of view because of an increased risk of nitrate leaching. In an ideal production system, N supply would be synchronized with plant demand. On the other hand, cultural practices like tillage or cultivation may reduce the complexity of the soil food web. Thus, a decrease in the frequency and intensity of tillage may promote the conservation of predatory nematodes and contribute to improved farming system performance.

References
  • Ferris, H., T. Bongers, and R. G. M. de Goede. 2001. A framework for soil food web diagnostics: Extension of the nematode faunal analysis concept. Applied Soil Ecology 18: 13–29.
  • Ferris, H., R.C. Venette, H.R. van der Meulen, and S.S. Lau. 1998. Nitrogen mineralization by bacterial-feeding nematodes: Verification and measurement. Plant and Soil 203: 159–171.
  • Freckman, D.W., and C.H. Ettema. 1993. Assessing nematode communities in agroecosystems of varying human intervention. Agriculture, Ecosystems & Environment 45: 239–261.
  • Griffiths, B.S., K. Ritz, and R.E. Wheatley. 1994. Nematodes as indicators of enhanced microbiological activity in a Scottish organic farming system. Soil Use and Management. 10: 20–24.
  • Lavelle, P., and A.V. Spain. 2001. Soil ecology. Kluwer Academic Publishers, Boston, MA.
  • Nahar, M.S., P.S. Grewal, S.A. Miller, D. Stinner, B.R. Stinner, M.D. Kleinhenz, A. Wszelaki, and D. Doohan. 2006. Differential effects of raw and composted manure on nematode community, and its indicative value for soil microbial, physical and chemical properties. Applied Soil Ecology 34: 140–151.
  • Neher, D.A. 1999. Soil community composition and ecosystem processes: Comparing agricultural systems with natural ecosystems. Agroforestry Systems 45: 159–185.
  • Peterson, H., and M. Luxton. 1982. A comparative analysis of soil fauna populations and their role in decomposition processes. Oikos 39: 287–388.
  • Stirling, G.R. 1991. Biological control of plant parasitic nematodes. CAB International, Wallingford, U.K.
Further Reading
  • Ferris, H. 1998. The role of nematodes in soil fertility [Online]. NEMAPLEX: the Nematode–plant expert information system. A virtual encyclopedia on soil and plant nematodes. University of California. Available at: http://plpnemweb.ucdavis.edu/Nemaplex/Ecology/fertil.htm (verified 4 April 2011).
  • Guerena, M. 2006. Nematodes: Alternative controls [Online]. ATTRA publication #IP287. Available at: http://attra.ncat.org/attra-pub/nematode.html (verified 4 April 2011).
  • Ugarte, C., M. Wander, and E. Zaborski. 2006. So you want to manage soil food webs? Focus on nematodes [Online]. New Agricultural Network, Vol. 3 No. 8. Available at: iwww.ipm.msu.edu/new-ag/issues06/7-26.htm (verified 4 April 2011).

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 4495

Certification of Organic Farming Systems

ven, 2012/08/31 - 12:54

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic T869

Organic Seed Growers Conference 2012: Selected Live Broadcasts

jeu, 2012/08/30 - 17:35

eOrganic and the Organic Seed Alliance brings you selected live broadcasts from the Organic Seed Growers Conference in Port Townsend, WA. This conference brings together hundreds of farmers, seed production and distribution companies, researchers, plant breeders, pathologists, and university extension in two days of informative presentations, panel discussions, and networking events.

Visit the Organic Seed Alliance's website to learn more about the Organic Seed Grower's Conference »

Find other upcoming and recorded eOrganic webinars »

Recordings and Slides

 

Introduction to On-Farm Plant Breeding Workshop.

An increasing number of farmers are starting to breed new varieties and reselect older varieties for their farms. This presentation will introduce you to the steps needed to create new crop varieties on your farm with little or no hand-pollination or specialized tools. Presenter: John Navazio, Organic Seed Alliance and Washington State University

 

Organic Wheat Breeding Workshop

With the explosion of local organic grains, mills and bakeries, organic farmers are looking for wheat varieties that thrive in their systems. This workshop will take you through the process of creating your own wheat variety and describe some of the current organic what breeding projects. Presenters: Stephen Jones, Washington State University; Richard Little, University of Nebraska-Lincoln; Dean Spaner, University of Alberta

  • Winter Wheat Breeding Basics. Richard Little - Video | Handout
  • Organic Wheat Breeding and Agronomy Research. Dean Spaner - Video | Handout

 

Breeding Peas, Sweet Corn, Broccoli, Winter Squash and Carrots as part of NOVIC

NOVIC is a national project to breed new vegetable varieties for organic agriculture. You will learn from the panelists about the techniques they are using to breed new organically adapted varieties of peas, sweet corn, broccoli, squash, and carrots. Presenters: Jim Myers, Oregon State University; Michael Mazourek, Cornell University; William Tracy, University of Wisconsin-Madison; John Navazio, Organic Seed Alliance and Washington State University; Laurie McKenzie, Oregon State University; Adrienne Shelton, University of Wisconsin.

 

Organic Corn Breeding Workshop

King corn is grown on more acres than any other crop. What is being done to breed corn for organic systems, and how can you take part? This workshop will describe the process of breeding corn for organic agriculture and some of the current organic corn breeding projects. Presenters: Frank Kutka, NPSAS Farm Breeding Club; William Tracy, University of Wisconsin-Madison.

  • Corn Reproduction: Inbreds and Hybrids. William Tracy
  • Sweet Corn: Organic Breeding Considerations. William Tracy
  • Breeding High Quality Corn for Sustainable and Organic Farmers. Walter Goldstein

 

Breeding for Nutrition Workshop

This broadcast was repeated as an eOrganic webinar on March 23, 2012. Please find the handout for the webinar here.

Organic eaters want nutritious food, but some modern breeding programs may be increasing yields at the cost of nutrition. Learn about breeding programs working with classical breeding methods (non-gmo) to breed nutritionally superior crops.

  • Prospects and Challenges for Plant Breeders. Philipp Simon - Video | Handout
  • Breeding Tomatoes for Increased Flavonoids. Jim Myers - Video | Handout
  • Breeding Corn for Nutritional Value. Walter Goldstein - Video | Handout
  • Full version with all 3 presentations and discussion - Video | Handout

 

Breeding for Positive Microbial Interactions Workshop

We know that many beneficial soil microorganisms provide plants with access to nutrients, improve water uptake and even have the potential to suppress certain soil borne diseases. The ability to breed plants to optimize their interaction with the soil microbiology holds great potential to enhance organic farming systems. Hear about the latest studies in this important and expanding field of science.

  • Wheat Varietal Selection and Annual Versus Perennial Growth Habit Impact Soil Microbes and Apple Replant Disease Suppression. Lori Hoagland -Video | Handout
  • Linking Hairy Vetch Germplasm Diversity to Traits Facilitating Improved Nitrogen Fixation. Jude Maul -Video | Handout
  • Breeding Corn for Positive Soil Microbial Interactions. Walter Goldstein -Video | Handout

organic seed growers conference 2012

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 7461

Webinars by eOrganic

mar, 2012/08/21 - 20:01
imageLive and archived webinars on organic farming and research.

Learn the latest in organic farming practices and research by attending or watching an eOrganic Webinar. Sign up for upcoming Webinars to watch slides, listen to the presenter, and type in questions during the live event. To receive notices about upcoming Webinars, and when we post the archived sessions, sign up for the eOrganic newsletter.

Learne how to earn CCA credits for eOrganic webinar recordings

 

Archived Webinars Link Presenters Date CCA
Credits Sourcing Organic Seed Just Got Easier: An Introduction to Organic Seed Finder Watch Chet Boruff, AOSCA, Kristina Hubbard, Organic Seed Alliance August 21, 2012   Your Organic Dairy Herd Health Toolbox Watch Dr. Hubert Karreman, Penn Dutch Cow Care July 16, 2012   International Organic Fruit Symposium Recordings coming soon
various June 19 and 21, 2012   Breeding and Genetics: Considerations for Organic Dairy Farms Watch
Brad Heins, University of Minnesota June 19, 2012   Organic Weed Management on Livestock Pastures Watch
Sid Bosworth, University of Vermont 5/15/12   Live Broadcast from Fly Management on Your Organic Dairy Workshop Watch Roger Moon, University of Minnesota; J Keith Waldron, Cornell; Wes Watson, North Carolina State University 4/19/12   NRCS EQIP Technical and Financial Support for Conservation on Organic Farms Webinar Watch Sarah Brown, Oregon Tilth 3/29/12   Organic Seed Breeding for Nutrition Watch
Philipp Simon, Walter Goldstein, Jim Myers, Micaela Colley 3/23/12   Cover Crops for Disease Suppression Watch
Alex Stone, Oregon State University 3/20/12   Fire Blight Control in Organic Pome Fruit Systems Under the Proposed Non-antibiotic Standard Watch
Ken Johnson, Oregon State University, Rachel Elkins, UC Cooperative Extension 3/13/12   The Role of Cover Crops in Organic Transition Strategies Watch Brian McSpadden Gardener, The Ohio State University 3/6/12   Optimizing the Benefits of Hairy Vetch in Organic Production Watch John Teasdale, USDA-ARS Sustainable Agricultural Systems Lab, Beltsville, MD 2/28/12   Stink Bug Management with Trap Crops Watch
Russell Mizell, University of Florida 2/21/12   Veggie Compass: Whole Farm Profit Management
Watch Erin Silva and Rebecca Claypool, University of Wisconsin-Madison 2/14/12   Cultivation and Seedbank Management for Improved Weed Control Watch Eric Gallandt, University of Maine 2/7/12   Participatory On-farm Research: Beyond the Randomized Complete Block Design Watch Sieglinde Snapp, Michigan State University 1/31/12   The OrganicA Project: Current Research on Organic Production of Ginger Gold, Honeycrisp, Zestar!, Macoun, and Liberty Apples Watch Lorraine Berkett, University of Vermont 1/24/12   The Organic Seed Grower's Conference, Port Townsend Washington: Selected Live Broadcasts Watch various 1/20/12 and 1/21/12   Ecological Farm Design for Pest Management In Organic Vegetable Production: Successes and Challenges on Two Farms Watch Helen Atthowe, Doug O'Brien 1/18/12   Carolina Organic Commodities and Livestock Conference: Selected Live Broadcasts Watch various 1/12/12 and 1/13/12   Why Eat Organic: Live Broadcast from the Illinois Specialty Crops, Agritourism and Organic Conference Watch Jim Riddle, University of Minnesota 1/12/12   Reduced Tillage in Organic Vegetable Production: Successes, Challenges, and New Directions Watch Helen Atthowe, Biodesign Farm, Consultant 12/13/11   Microbial Food Safety Issues of Organic Foods Watch Francisco Diez-Gonzalez, University of Minnesota 12/6/11   Starting Up Small-Scale Organic Hops Production Watch Rob Sirrine, Michigan State University, Brian Tennis, Michigan Hop Alliance 11/15/11   Dryland Organic Agriculture Symposium from the Washington Tilth Conference 2011 Watch Various speakers, morning and afternoon sessions. 11/11/11   Tracking Your Produce--For Your Business and Health Watch Collen Collier Bess, Michigan Dept of Agriculture 11/8/11   Healthy Soils for a Healthy Organic Dairy Farm -- Broadcast from 2011 NOFA-NY Organic Dairy Conference Watch Heather Darby, University of Vermont, Cindy Daley, University of California, Chico 11/4/11   Root Media and Fertility Management for Organic Transplants Watch John Biernbaum, Michigan State University 11/1/11   Plan for Marketing Your Organic Products Watch Susan Smalley, Michigan State University 10/25/11   How to Breed for Organic Production Systems Watch Jim Myers, Oregon State University 10/18/11   Flooding and Organic Certification Watch Jim Riddle, University of Minnesota 10/13/11   Stockpiling Forages to Extend the Grazing Season on Your Organic Dairy Watch
Laura Paine, Wisconsin Department of Agriculture, Trade and Consumer Protection 7/28/11   Fly Management in the Organic Dairy Pasture Watch Donald Rutz, Keith Waldron, New York State IPM Program 7/6/11   Using Small Grains as Forages on Your Organic Dairy Watch Heather Darby, University of Vermont Extension 4/14/11   Third Party Audits for Small and Medium Sized Meat Processors Watch Jim Riddle, Joe McCommons, and the Quality Control Manager of Lorentz Meats 4/5/11   A Novel Strategy for Soil-borne Disease Management: Anaerobic Soil Disinfestation (ASD) Watch Joji Muramoto, Carol Shennan, David Butler, Maren Mochizuki, Erin Rosskopf 3/30/11 Earn credit Integrated Pest Management in Organic Field Crops Watch Eileen Cullen. Robin Mittenthal, University of Wisconsin, Christine Mason, Standard Process Farm 3/29/11 Earn credit The Evolution, Status, and Future of Organic No-Till in the Northeast US Watch Bill Curran, Penn State, Steven Mirsky, USDA, Bill Mason, Mason's Heritage Farms 3/22/11   USDA ERS 2011 Organic Farming Systems Conference Webinars Watch various 3/16/11   Local Dirt: Beyond Marketing. Find Buyers, Sell Online, Source & Buy Product…Yourself Watch Heather Hilleren, Kassie Rizzo, Local Dirt 3/15/11   GMO Contamination: What's an Organic Farmer to Do? Watch Jim Riddle, University of Minnesota 3/9/11   North Carolina's Statewide Initiative for Building a Local Food Economy Watch Nancy Creamer, Teisha Wymore, North Carolina State University 3/1/11   Grafting for Disease Management in Organic Tomato Production Watch Frank Louws North Carolina State University Cary Rivard, Kansas State University 2/22/11   Shades of Green Dairy Farm Calculator Watch Charles Benbrook, The Organic Center 2/1/11   Greenhouse Gas Emissions Associated with Dairy Farming Systems Watch Tom Richard, Gustavo Camargo, Penn State 1/25/11   Assessing Nitrogen Contribution and Rhizobia Diversity Associated with Winter Legume Cover Crops in Organic Systems Watch Julie Grossman, North Carolina State University 12/14/10   Using Winter Killed Cover Crops to Facilitate Organic No-till Planting of Early Spring Vegetables Watch Mike Snow, Farm Manager, Accokeek Ecosystem Farm; Charlie White, Penn State 12/7/10   Using Cover Crops to Suppress Weeds in Northeast US Farming systems Watch William Curran, Matthew Ryan, Penn State 12/2/10   Transitioning Organic Dairy Cows off and on Pasture Watch Rick Kersbergen, University of Maine 11/23/10   Greenhouse Gases and Agriculture: Where does Organic Farming fit? Watch David Granatstein, Lynne Carpenter-Boggs, Washington State University, Dave Huggins 11/15/10   Impact of Grain Farming Methods on Climate Change Watch Michel Cavigelli, USDA, Beltsville MD 11/12/10   Setting up a Grazing System on Your Organic Dairy Farm Watch Sarah Flack, Sarah Flack Consulting, Cindy Daley, California State University, Chico 10/1/10   Maximizing Dry Matter Intake on Your Organic Dairy Farm Watch Karen Hoffman, USDA-NRCS 9/16/10   How to Calculate Pasture Dry Matter Intake on Your Organic Dairy Farm Watch Sarah Flack, Sarah Flack Consulting 8/20/10   Late Blight Control in Your Organic Garden Watch Meg McGrath, Cornell 7/21/10   Late Blight Control on Organic Farms Watch Meg McGrath, Cornell, Sally Miller, Ohio State 7/1/10   Increasing Plant and Soil Biodiversity on Organic Farmscapes Watch Louise Jackson, University of California-Davis 5/4/10   Cover Crop Selection Watch Jude Maul, USDA ARS 4/27/10   The Economics of Organic Dairy Farming in New England Watch Bob Parsons, University of Vermont 4/13/10   Estimating Plant-Available Nitrogen Contribution from Cover Crops Watch Nick Andrews, Dan Sullivan, Oregon State 4/13/10   Planning for Flexibility in Effective Crop Rotations Watch Chuck Mohler, Cornell 4/6/10   Using NRCS Conservation Practices and Programs to Transition to Organic Watch David Lamm, USDA NRCS 3/30/10   Planning Your Organic Farm for Profit Watch Richard Wiswall, Cate Farm 3/22/10   A Look at the Newly Released Organic Pasture Rule Watch Kerry Smith, USDA, AMS, National Organic Program 3/17/10   Organic Blueberry Production Watch Bernadine Strik, Handell Larco, Oregon State University, David Bryla, USDA 3/9/10   High Tunnel Production and Low Cost Tunnel Construction Watch Tim Coolong, University of Kentucky 3/2/10   Getting EQIPed: USDA Conservation Programs for Organic and Transistioning Farmers Watch Jim Riddle, University of Minnesota 2/23/10   Organic Certification of Research Sites and Facilities Watch Jim Riddle, University of Minnesota 2/9/10   Grafting Tomatoes for Organic Open Field and High Tunnel Production Watch David Francis, Ohio State 2/2/10   Undercover Nutrient Investigation: The Effects of Mulch on Nutrients for Blueberry Watch Dan Sullivan, Ryan Costello, Luis Valenzuela, Oregon State 1/26/10   ABCs of Organic Certification Watch Jim Riddle, University of Minnesota 1/19/10   Organic Farming Financial Benchmarks Watch Dale Nordquist, University of Minnesota 1/12/10  

 

This is an eOrganic article and was reviewed for compliance with National Organic Program regulations by members of the eOrganic community. Always check with your organic certification agency before adopting new practices or using new materials. For more information, refer to eOrganic's articles on organic certification.

eOrganic 4942

Pages