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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 3 heures 10 min

Planning for Ecological Weed Management in Organic Vegetable Production

lun, 2012/10/15 - 14:58

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 T1047

Webinar: Can we talk? Improving weed management communication between organic farmers and Extension

lun, 2012/10/15 - 14:49

Join eOrganic for an organic weed management research webinar on November 13 at 2PM Eastern Time (1PM Central, 12PM Mountain, 11AM Pacific Time. The webinar is free and open to the public, and the topic is directed at Extension, government agency staff and researchers. Space is limited and advance registration is required.
Register now at: https://www1.gotomeeting.com/register/701945704

About the Webinar

Understanding how farmers make decisions, not just what decisions they make, can improve our communication with farmers and our ability to provide relevant information that builds upon their existing knowledge, perceptions, and values. Sarah Zwickle and Marleen Riemens will discuss their research on organic farmers’ weed management beliefs, perceptions and behaviors, and how it can contribute to our research and extension efforts with organic farmers.

About the Presenters

Sarah Zwickle is a research assistant in the School of Environment and Natural Resources at the Ohio State University. Her master’s research on the weed management decision-making process of organic farmers has served as the foundation for the extension and outreach efforts of the OREI project “Mental Models and Participatory Research to Redesign Extension Programming for Organic Weed Management”.

Marleen Riemens is a weed scientist at the Netherlands’ Wageningen University & Research Centre. For the past few years, her research has extended its scope to include the relationship between weed pressure on organic farms and organic farmers’ weed management behaviors and beliefs.

Find all upcoming and archived eOrganic webinars at http://www.extension.org/pages/25242

System Requirements

PC-based attendees
Required: Windows® 7, Vista, XP or 2003 Server
Macintosh®-based attendees
Required: Mac OS® X 10.5 or newer

Java needs to be installed and working on your computer to join the webinar. If you have concerns, please test your Java at http://java.com/en/download/testjava.jsp prior to joining the webinar. If you are running Mac OS X 10.5 with Safari, please be sure to test your Java. If it isn't working, please try Firefox (http://www.mozilla.com) or Chrome (http://www.google.com/chrome).

PC-based attendees
Required: Windows® 7, Vista, XP or 2003 Server
Macintosh®-based attendees
Required: Mac OS® X 10.5 or newer
Mobile attendees
Required: iPhone®, iPad®, Android™ phone or Android tablet

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 8277

Using the eOrganic Organic Seed Production Tutorials Webinar

lun, 2012/10/15 - 13:58

Join eOrganic for a webinar on the Organic Seed Production tutorials created by the Organic Seed Alliance. The webinar is free and open to the public, and advance registration is required. The webinar will take place on Friday, November 16 at 1PM Eastern Time (12PM Central, 11AM Mountain, 10AM Pacific Time).

Register at https://www1.gotomeeting.com/register/675611681

About the Webinar

Webinar participants can expect to learn about the Organic Seed Production tutorials, and how to most effectively access in-depth organic seed production information available through this new eOrganic course created by Organic Seed Alliance. Find the tutorials at http://campus.extension.org/course/view.php?id=377.

About the Presenter

Jared Zystro is the California research and education specialist for Organic Seed  Alliance. Jared designed the eOrganic Organic Seed Production Tutorials.

About eOrganic

eOrganic is the Organic Agriculture Community of Practice at eXtension.org. Our website  at http:www.extension.org/organic_production contains articles, videos, and webinars for farmers, ranchers, agricultural professionals, certifiers, researchers and educators seeking reliable information on organic agriculture, published research results, farmer experiences, and certification. The content is collaboratively authored and reviewed by our community of University researchers and Extension personnel, agricultural professionals, farmers, and certifiers with experience and expertise in organic agriculture.

Find all upcoming and archived eOrganic webinars at http://www.extension.org/pages/25242

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 8263

Going Organic Has Challenges

mar, 2012/09/25 - 14:43

Released September 23, 2012

BISMARCK,N.D. - Despite a recent Stanford University study disputing extra health benefits of organic farming, organic producers do not expect their industry to be affected.

Instead, Brad Brummond president of the North Dakota Organic Advisory Board, said demand seems to be increasing, but the number of producers seems to be decreasing.

“Our biggest challenge is to replace them,” he said.

--continued on the Bismarck Tribune, http://bismarcktribune.com/business/local/going-organic-has-challenges/a...

Integrated Pest Management in Organic Field Crops Webinar

mar, 2012/09/25 - 11:42

 

 

About the webinar

In this webinar, recorded on, March 29, 2011, a farmer,  Christine Mason, shares her approach to minimizing insect pest impact in organic farming systems, while University of Wisconsin researchers Eileen Cullen and Robin Mittenthal weave in results from projects specifically designed to advance the IPM paradigm for organic agriculture.

Slides from the webinar as a pdf file: http://cop.extension.org/mediawiki/files/f/f3/IPMWebinar.pdf

Resources mentioned in the webinar:

UC Davis IPM Degree Days calculator:  http://www.ipm.ucdavis.edu/WEATHER/ddretrieve.html

University of Wisconsin degree day calculator: http://www.soils.wisc.edu/uwex_agwx/thermal_models/degree_days

Trap Cropping and Insect Control  on Cornell University's Resource Guide for Organic Insect and Disease Management

About the Presenters

Christine Mason is Farm Manager at Standard Process certified-organic farm, Palmyra, Wisconsin, Secretary of the Wisconsin Organic Advisory Council, and a Certified Crop Advisor. Christine and her family are the fifth generation on their family farm in Palymyra, WI growing organic corn, soybeans, wheat and forages.

Eileen Cullen is Associate Professor at University of Wisconsin-Madison Entomology Department. She is also UW-Extension State Specialist for field and forage crop entomology focusing on integrated pest management (IPM).

Robin Mittenthal has worked on organic farms, taught high school science, and is now completing his doctorate in entomology at the University of Wisconsin with a focus on connections between soil fertility, plant health, and insect responses.

About eOrganic

eOrganic is the Organic Agriculture Community of Practice at eXtension.org. Our website  at http:www.extension.org/organic_production contains articles, videos, and webinars for farmers, ranchers, agricultural professionals, certifiers, researchers and educators seeking reliable information on organic agriculture, published research results, farmer experiences, and certification. The content is collaboratively authored and reviewed by our community of University researchers and Extension personnel, agricultural professionals, farmers, and certifiers with experience and expertise in organic agriculture.

Find all upcoming and archived eOrganic webinars at http://www.extension.org/pages/25242

 

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 5810

Weed Management in Organic Systems

mer, 2012/09/12 - 17:03

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 T877

Organic Vegetable Production Systems

mer, 2012/09/12 - 17:03

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

Organic Vegetable Production Systems, Weed Management in Organic Systems

mer, 2012/09/12 - 17:03

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 T879,877

Yellow Nutsedge (Cyperus esculentus) in Greater Depth

mer, 2012/09/12 - 17:00

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Introduction

Yellow nutsedge (Cyperus esculentus) is a major weed of vegetable and row crops in temperate and tropical regions around the world. Native to the Americas, yellow nutsedge is especially aggressive in irrigated crops that are maintained at high soil moisture levels (Ransom et al., 2009), and is considered a major weed of vegetables, corn, cotton, and peanuts in the southern United States (Holm et al., 1991; Webster, 2006).

Yellow nutsedge is a grass-like weed in the sedge family (Cyperaceae) with top growth 8–30 inches tall (Fig. 1a), and an extensive underground network of basal bulbs, roots, thin fibrous rhizomes, and tubers 0.4–0.8 inch long borne singly at the tips of rhizomes (Uva et al., 1997; Holm et al., 1991). The leaves are mostly basal, bright green to yellow-green, 0.1–0.35 inch wide with a prominent midrib, and tapering gradually toward the tips. Leaves may be as long as, or longer than, the culm (stem), which is triangular in cross section, and bears the inflorescence (flower head). The inflorescence is yellow-brown, golden, or straw colored, and consists of an umbel of spikes borne on stalks of unequal length (1–3 inches), subtended by leaflike bracts as long or longer than the spikes (Bryson and DeFelice, 2009) (Fig. 1b).

Yellow nutsedge in bloom.
Figure 1. (a) Yellow nutsedge in bloom. (b) Closeup of yellow nutsedge inflorescence, showing umbel of spikes of variable length, and leaflike bracts longer than the spikes. Photo credits: (a) Howard F. Schwartz, Colorado State University, Bugwood.org; (b) Mark Schonbeck, Virginia Association for Biological Farming.

Biology

Tuber dormancy in yellow nutsedge is broken by chilling at 35–50 °F for several weeks. Sprouting begins as soil temperatures rise above 55 °F (Holm et al., 1991; Stoller and Wax, 1973), and is promoted by tillage. Washing tubers has enhanced germination by 8-fold (Tumbleson and Kommendahl, 1961), which suggests that a water soluble inhibitor contributes to tuber dormancy, and that percolation of moisture through the soil may play a role in promoting germination (Mohler and DiTommaso, unpublished). Although the majority of tubers either sprout or die during the first year after they are formed, some tubers located deeper in the soil profile can remain dormant and survive 2–4 years (Stoller and Wax, 1973; California Department of Food and Agriculture) .

Yellow nutsedge begins active growth in mid- to late spring (FIg. 2a). A determinate and fairly short rhizome emerges from the tuber and grows toward the soil surface. When the tip of the rhizome receives a light stimulus, a basal bulb forms about 0.4–0.8 inch behind the tip (Stoller et al., 1972). The shoot, consisting of a cluster of basal leaves, arises from this bulb. In the field, basal bulbs generally form within 0.4–1.6 inches of the soil surface (Fig. 2b), regardless of the depth of the tubers from which they arise (Mulligan and Junkins,1976; Stoller et al., 1972; Uva et al., 1997). A fibrous root system develops from basal bulbs and rhizomes.

As the initial plant develops, it remains attached to the mother tuber for up to 12 weeks (Stoller et al., 1972). Within four weeks after initial shoot emergence, new indeterminate rhizomes emerge from the basal bulb, grow 1–20 inches laterally, and form new basal bulbs and daughter plants (Fig. 2b, Fig. 3). The cycle repeats several times, with new shoots continuing to develop through July in the central U.S. (Jordan-Molero and Stoller, 1978); thus, the weed spreads exponentially in the absence of competition or control measures.

Yellow nutsedge in a vegetable garden
Figure 2. Yellow nutsedge in a home garden in central Virginia, photographed in late May. (a) Growing with snap bean (b) Plants exhumed to show basal bulbs about 1–1.5 inches below the soil surface, with fibrous roots growing down into the soil, and white rhizomes extending laterally to establish daughter plants. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

Yellow nutsedge roots and rhizomes.
Figure 3. Yellow nutsedge specimens collected in late May in Virginia, about a month after emergence: (a) still attached to the mother tuber; (b) sending out several rhizomes, two of which have already initiated daughter plants. Photo credits: Mark Schonbeck, Virginia Association for Biological Farming.

In temperate zone populations of yellow nutsedge, shortening daylength in late summer triggers flowering and tuber production. When daylength decreases to about 14 hours, rhizome tips begin to form tubers rather than new daughter plants. This has been reported to occur in late July in Ohio (Cardina et al., 2012), near the beginning of August in Illinois (Jordan-Molero and Stoller, 1978), and mid to late August in Canada (Mulligan and Junkins, 1976). While top growth slows, prolific tuber production continues until killing frost. Tubers are formed mostly throughout the top 6–10 inches of the soil profile, although a few may form as deep as 18 inches (Tumbleson and Kommedahl, 1961). The mature, starchy tubers contain about 40-60% dry matter.

Yellow nutsedge thrives in moist to wet conditions (Fig. 4), and is highly tolerant to flooding. It can be incredibly prolific in temperate climates and high-moisture soils. A single tuber has been observed to give rise to 1,900 shoots and 6,900 tubers within one year in Minnesota (Tumbleson and Kommedahl, 1961), and 1,700–3,000 shoots and 19–20 thousand tubers in irrigated fields in Oregon (Ransom et al., 2009), forming a dense patch about 6 feet across. Tuber dry weight reached an equivalent of about 4 tons per acre. In the Oregon study, yellow nutsedge spread several fold more slowly when the soil was allowed to dry out between infrequent irrigations of 1.0–1.4 inches as recommended for sugarbeet and wheat, than when the soil was maintained near field capacity through frequent small (0.3 inch) irrigations as recommended for onion production. Yellow nutsedge spreads more slowly in very hot, sunny climates such as the low desert of southeastern California, where a single tuber gave rise to about 50 tubers after one growing season (Wang et al., 2008).

Yellow nutsedge response to soil drainage
Figure 4. A dense stand of yellow nutsedge infests the lowest and wettest part of this field (foreground), with much lighter populations in the better-drained parts of the field (background). Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Yellow nutsedge tubers are killed by exposure for 1–2 days to temperatures of 113–122 °F (Webster, 2003) or below 20 °F, although cold hardening may enhance freeze tolerance by a few degrees (Stoller and Wax, 1973). In Illinois, most tubers within 2 inches of the soil surface are winterkilled but the weed readily emerges from tubers located 4–8 inches deep, where temperatures extremes are buffered. Yellow nutsedge has successfully spread into southeastern Canada, where snow cover may further protect tubers from winterkill. Desiccation or exposure to direct sunlight accentuates thermal stresses, and tubers brought to the surface by tillage rapidly lose viability during dry weather (Tumbleson and Kommedahl, 1961). Intact yellow nutsedge is quite tolerant to drought because of its extensive fibrous root system (Holm et al., 1991).

Attempts to eradicate yellow nutsedge by soil solarization in Georgia have been thwarted by survival of tubers deeper in the soil profile (Webster, 2003). However, solarization has successfully controlled this weed in the low desert of California (Wang et al., 2008).

Yellow nutsedge tolerates moderate (20–30%) shade with little decrease in growth or tuber production, whereas 60–80% shade reduces total biomass by more than half, and tuber biomass by 75% or more (Jordan-Molero and Stoller, 1978; Keeley and Thullen, 1978; Santos et al., 1997). Although the weed compensates for shade by growing taller, and can form some tubers even under 94% shade, crop competition for light is recognized as an important tactic that can enhance the efficacy of cultivation (Keeley and Thullen, 1978).

Yellow nutsedge can form viable seeds by cross-pollination between different clones (populations of genetically identical plants arising from vegetative reproduction) (Fig. 5). As many as 1,500 viable seeds per plant have been reported, yet yellow nutsedge seedlings are rarely observed in the field (Holm et al., 1991; Mohler and DiTommaso, unpublished). The seedlings are extremely sensitive to desiccation, and can establish successfully only when the soil surface remains continuously moist (Lapham and Drennan, 1980). Although sexual reproduction is not a significant means of propagation, its occasional occurrence allows for genetic recombination and adaptation. Morphological and ecophysiological differences among yellow nutsedge populations have been documented (Mulligan and Junkins, 1976), and an unusually vigorous biotype has been reported in California (Tayyar et al., 2003).

Yellow nutsedge seeds
Figure 5. Yellow nutsedge seeds. Photo credit: Ken Chamberlain, The Ohio State University, Bugwood.org.

Impacts on Crop Production

Yellow nutsedge competes severely against many crops for moisture and nutrients (Fig. 6), and against low-growing crops for light. Its sharp pointed rhizomes can penetrate the edible portion of root crops, rendering them unmarketable; severe crop losses can occur in potato (Holm et al., 1991). Crop competition from tall crops like corn and tomato, and heavy canopy formers like potato can suppress nutsedge that emerges several weeks after crop planting, whereas slow-starting crops like onion and cotton, and warm season crops planted early when the soil is still cool, are especially prone to nutsedge competition. Potato growers in Ontario have noted that the variety ‘Kennebec,’ which is characterized by vigorous top growth, reduces yellow nutsedge populations in the following year, compared to less vigorous varieties such as ‘Katahdan’ and ‘Irish Cobbler’ (Mulligan and Jenkins, 1976).

Yellow nutsedge infestations
Figure 6. (a) A dense patch of yellow nutsedge in onion. (b) Heavy infestation of yellow nutsedge in cotton has virtually hidden the crop from view. Figure credits: (a) Howard F. Schwartz, Colorado State University, Bugwood.org; (b) John D. Byrd, Mississippi State University, Bugwood.org.

Allelopathic substances from yellow nutsedge have reduced growth of corn, soybean, and sweet potato in greenhouse trials (Drost and Doll, 1980; Peterson and Harrison, 1995). Although allelopathy has been cited as a factor in corn and soybean yield losses to yellow nutsedge (Cardina et al., 2012), more research is needed to verify this in field conditions.

A few crops may suppress yellow nutsedge through allelopathy. In field studies in South Carolina, sweet potato reduced yellow nutsedge biomass production by 90% through competition and allelopathy, without itself being hindered by the weed (Harrison and Peterson, 1991). Rye straw mulch and root residues in the soil left by roll-killing a winter rye cover crop can reduce yellow nutsedge growth (Mohler and DiTommaso, unpublished). In greenhouse trials, wild radish residues mixed into the potting soil reduced yellow nutsedge competitiveness against tomato and pepper (Norsworthy and Meehan, 2005a). However, yellow nutsedge tubers showed little sensitivity to several isothiocyanate allelochemicals released by decaying Brassica residues (Norsworthy and Meehan, 2005b).

Emerging shoots of yellow nutsedge readily penetrate organic mulches and the lighter grades of spun-fiber mulches. Other synthetic mulches, such as black polyethylene mulch can slow emergence and spread of yellow nutsedge by about half (Webster 2005a and 2005b); however, some plants pierce the mulch, which can complicate end-of-season mulch removal. Because shoot tips open in response to light, yellow nutsedge emerging under clear polyethylene may become trapped and heat-killed under the film, which weakens the mother tuber (Chase et al., 1998).

Yellow nutsedge did not show a strong response to fertilizer nitrogen (N) rate in Oregon (Ransom et al., 2009), and does not appear to compete more aggressively against crops at high levels of available nutrients in the Northeast (Mohler and DiTommaso, unpublished). However, in a Florida field trial, yellow nutsedge competed more aggressively against tomato at higher N application rates (Morales-Payan et al., 1997).

Yellow nutsedge thrives in moderately warm conditions with ample moisture. It is most troublesome in irrigated crops such as onion, which are maintained at high soil water potential (Ransom et al., 2009). One organic farmer in Arkansas has observed that yellow nutsedge is most abundant in fields with compacted soil and poor drainage, and becomes less so after organic management has improved soil drainage and soil quality (Josh Hardin, personal communication, 2009). Compared to its close relative purple nutsedge, yellow nutsedge was much easier to control and had far less impact on yields of irrigated broccoli in field trials in the low desert of southeastern California (Wang et al., 2008, Wang et al., 2009).

Management Strategies and Tactics for Organic Production

Timely tillage and cultivation during the initial flush of yellow nutsedge emergence in late spring can significantly weaken the weed. In experiments on Illinois populations, separating young plants (about 1 inch of shoot emerged above ground) from the mother tuber substantially reduced subsequent growth and spread (Stoller et al., 1972). Tubers that were germinated for 6 days at 77 °F in darkness expended over 60% of their dry weight, protein, carbohydrate, and oils to develop the sprout. Many tubers sprouted again after the initial sprout was removed, but the second sprout was much weaker (Stoller et al., 1972). Thus, the weed is most vulnerable to tillage in late spring, after tubers have sprouted but before the emerging shoot has begun to build up new reserves or initiate daughter plants. Tillage at this time and again in late summer (to disrupt formation of new tubers), combined with crop competition, have been found to control yellow nutsedge fairly well in the Northeast (Mohler and DiTommaso, unpublished).

One tillage operation will not control yellow nutsedge, and regrowth needs to be removed before the weed begins to rebuild underground reserves. Repeat cultivation before re-emerging shoots reach the 6-leaf stage (Russ and Burgess, 2009).

Yellow nutsedge that emerges in crops can be managed with cultivation. Farmers have had excellent results using a cultivator equipped with beet knives set about 1.5–2 inches below the surface to undercut the new growth and separate it from the tubers.

Repeated tillage alone may not be sufficient to eradicate a severe yellow nutsedge infestation. Two years of mechanical fallow throughout the frost-free period have reduced populations by 80–90% (Mulligan and Junkins, 1976; Tumbleson and Kommendahl, 1961); however the costs to soil quality and the production foregone through extended fallow make this approach impractical for most farms. Sustainable management of this weed requires an integrated strategy that subjects the weed to multiple stresses, especially during the critical times of initial emergence (mid–late spring) and tuber set (late summer). Key components include:

  • Till or cultivate at critical times, working 1.5–2 inches deep to undercut basal bulbs when practical.
  • Choose competitive crops and varieties, and manage crops to produce at least 80% shade between rows at ground level after the last cultivation.
  • Plan crop rotations and schedule plantings and field operations to allow either cultivation or effective crop competition during the period of active nutsedge growth, usually mid–late spring through late summer.
  • Follow spring vegetables with fast-growing, high-biomass summer cover crops to shade out nutsedge.
  • Avoid overwatering and overfertilizing—rotate nutsedge-infested fields to crops that do not require high soil moisture levels or high levels of readily available N.
  • Remedy soil drainage problems—chisel plow to break hardpan if necessary, grow deep rooted cover crops, and build soil organic matter.
  • Avoid synthetic mulches when moderate to high populations of yellow nutsedge are present.

Consider allelopathy from crops like rye and sweet potato, and soil solarization, as additional "little hammers" against yellow nutsedge, but do not rely on these tactics alone to control the weed.

Consider using livestock to reduce a heavy infestation. The tubers have a mild, sweet flavor that is especially attractive to pigs, which have been reported to consume most of the tubers in an infested area within a few days (California Department of Food and Agriculture). Poultry or weeder geese can also be effective. For food safety and to comply with USDA Organic Standards, be sure to remove livestock from the field and incorporate droppings into the soil at least 120 days prior to expected harvest of the next food crop. A good time to run livestock to clean up a weedy field is after a cash crop harvest and just before planting a cover crop.

Finally, farmers in the southern U.S. should obtain a definitive species identification of their nutsedge populations. Yellow and purple nutsedges often occur together in the South. Although they have similar life cycles, their responses to climate, environmental conditions, and control tactics can differ significantly; for example, purple nutsedge becomes far more aggressive in hot climates. Determine whether yellow, purple, or both nutsedges are present in order to fine-tune your management strategy.

References Cited
  • Bryson, C. T., and M. S. DeFelice. 2009. Weeds of the South. University of Georgia Press, Athens, GA.
  • California Department of Food and Agriculture. 2012. Cyperus genus. State of California. (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).
  • Drost, D. C., and J. D. Doll. 1980. The allelopathic effect of yellow nutsedge (Cyperus esculentus) on corn (Zea mays) and soybeans (Glycine max). Weed Science 28: 229–233. (Available online at: http://www.jstor.org/stable/4042988) (verified 10 Sept 2012).
  • Harrison, H. F., Jr., and J. K. Peterson. 1991. Evidence that sweet potato (Ipomoea batatas) is allelopathic to yellow nutsedge (Cyperus esculentus). Weed Science 39: 308–312. (Available online at: http://www.jstor.org/stable/4044934) (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.
  • Horak, M. J., J. S. Holt, and N. C. Ellstrand. 1987. Genetic variation in yellow nutsedge (Cyperus esculentus). Weed Science 35: 506-512. (Available online at: http://www.jstor.org/stable/4044520) (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).
  • Keeley, P. E., and R. J. Thullen. 1978. Light requirements of yellow nutsedge (Cyperus esculentus) and light interception by crops. Weed Science 26: 10–16. (Available online at: http://www.jstor.org/stable/4042682) (verified 10 Sept 2012).
  • Lapham, J., and D.S.H. Drennan. 1990. The fate of yellow nutsedge (Cyperus esculentus) seed and seedlings in soil. Weed Science 38: 125–128. (Available online at: http://www.jstor.org/stable/4045039) (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, 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).
  • Mulligan, G. A., and B. E. Junkins. 1976. The biology of Canadian weeds. 17. Cyperus esculentus L. Canadian Journal of Plant Science 56: 339–350.
  • Norsworthy, J. K., and J. T. Meehan, IV. 2005a. Wild radish-amended soil effects on yellow nutsedge (Cyperus esculentus) interference with tomato and bell pepper. Weed Science 53: 77–83. (Available online at: http://dx.doi.org/10.1614/WS-04-074R) (verified 10 Sept 2012).
  • Norsworthy, J. K., and J. T. Meehan, IV. 2005b. 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).
  • Cardina, J., C. Herms, T. Koch, and T. Webster. 2012. Yellow Nutsedge, Cyperus esculentus. Ohio Perennial and Biennial Weed Guide. The Ohio State University. (Available online at: http://www.oardc.ohio-state.edu/weedguide/singlerecord.asp?id=150) (verified ).
  • 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).
  • Ransom, C. V., C. A. Rice, and C. C. Shock. 2009. Yellow nutsedge (Cyperus esculentus) growth and reproduction in response to nitrogen and irrigation. Weed Science 57: 21–25. (Available online at: http://dx.doi.org/10.1614/WS-08-080.1) (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, T. A. Bewick, and D. G. Shilling. 1997. 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).
  • Stoller, E. W., D. P. Nema, and V. M. Bhan. 1972. Yellow nutsedge tuber germination and seedling development. Weed Science 20: 93–97. (Available online at: http://www.jstor.org/stable/4042040) (verified 10 Sept 2012).
  • Stoller, E. W., and L. M. Wax. 1973. Yellow nutsedge shoot emergence and tuber longevity. Weed Science 21: 76–81. (Available online at: http://www.jstor.org/stable/4042258) (verified 10 Sept 2012).
  • Tayyar, R. I., J.H.T. Nguyen, and J. S. Holt. 2003. Genetic and morphological analysis of two novel nutsedge biotypes from California. Weed Science 51: 731–739. (Available online at: http://dx.doi.org/10.1614/P2002-131) (verified 10 Sept 2012).
  • Tumbleson, M. E., and T. Kommedahl. 1961. Reproductive potential of Cyperus esculentus by tubers. Weeds 9: 646–653.
  • Uva, R. H., J. C. Neal, and J. M. DiTomaso, 1997. Weeds of the Northeast. Cornell University Press, Ithaca, NY.
  • 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).
  • Wang, G., M. E. McGiffen, Jr., E. J. Ogbuchiekwe, and L. Butler. 2009. Economic return of purple and yellow nutsedge management in vegetable production of southern California. Crop Protection 28: 319–326. (Available online at: http://dx.doi.org/10.1016/j.cropro.2008.11.011) (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).
  • 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 5135

Organic Dairy Production Systems

mer, 2012/09/12 - 16:59

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 T880

Synthetic Mulching Materials for Weed Management

mer, 2012/09/12 - 16:58

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Abstract

Synthetic mulches, especially black plastic film, effectively suppress most weeds, thereby reducing labor and other costs for weed control. Synthetic mulches can also improve growing conditions for the crop, and thereby improve crop competitiveness against weeds. Black plastic promotes crop growth and earliness by increasing soil temperature. Clear or translucent films warm the soil to a greater degree, but do not suppress weeds as effectively. Red, blue, white, and reflective mulches are occasionally used to enhance yields or repel insect pests in tomato, strawberry, and cucurbit crops. Red-on-black and white-on-black mulches combine yield and pest management benefits with the weed suppression of opaque films.

Concerns with labor and environmental costs of end-of-season disposal have led to the development of woven fabric mulches that can be used for 8–12 years, and of biodegradable paper mulches designed to suppress weeds for one season and decompose after harvest. This article discusses the properties, weed- and other management uses, advantages, and drawbacks of a variety of synthetic mulches.

Introduction

Vegetable producers have used plastic film mulches for at least 40 years to prevent weed growth near the crop, and to promote crop yields and earliness by modifying soil temperature and conserving soil moisture. Unlike organic mulches, both translucent and dark-colored opaque films speed soil warming, early crop growth, and ripening. Black polyethylene and other opaque films effectively suppress a wider spectrum of agricultural weeds than most organic mulches. Easy mechanical application, cost efficient weed control, and increased crop yields have led to widespread adoption of plastic mulch by organic and conventional vegetable farmers.

Disadvantages of plastic film mulch include the costs of mulch removal at the end of the season (required under USDA Organic Certification), petroleum consumption, waste generation, and the fact that plastic mulches do not build soil organic matter. A newer generation of synthetic mulches includes weed barrier or landscape fabric that lasts multiple seasons, several biodegradable mulches, and variously colored opaque or translucent films developed for specific crops or purposes.

Black Plastic Film

Black plastic film mulch, used in conjunction with in-row drip irrigation, is the weed management option of choice for many medium- to large-scale organic and conventional vegetable farms. The opaque film reduces germination of light-responsive weed seeds; shades out and physically blocks the emergence of most weeds; and can enhance crop growth by conserving soil moisture, promoting soil warming, and speeding nutrient mineralization from soil organic matter. The crop growth benefits contribute to weed management by enhancing the crop's ability to tolerate and compete with weeds.

The most economical and widely used synthetic mulches are plain or embossed black polyethylene films, which come in various widths (4 feet is most common) and thicknesses (0.8–1.25 mil), in rolls up to 2,500 feet or more in length. Usually, the mulch is applied by a tractor-drawn mulch layer that stretches the plastic evenly over crop rows or raised beds, anchoring the edges of the mulch with soil. Warm season crops like tomato, pepper, eggplant, melon, sweet potato, and okra are often grown on raised beds mulched with black plastic, and this system is sometimes used for early plantings of onion, lettuce, brassicas, and other cool-season crops as well (Fig. 1). Normally, drip lines are installed under the film (either on or below the soil surface) to deliver moisture and nutrients (liquid organic fertilizer) directly to the crop without watering and feeding between-row weeds.

Black plastic mulch
Figure 1. Black polyethylene mulch blocks weeds, reduces evaporative loss of soil moisture, and warms the soil, thereby promoting growth of early plantings of many vegetables, including lettuce (a) and tomato (b). Figure credits: (a) Becky Crouse, Marketing Manager, Potomac Vegetable Farms, Purcellville, VA; (b) Mark Schonbeck, Virginia Association for Biological Farming.

Black plastic mulch:

  • Can be mechanically applied to multi-acre fields with widely available equipment.
  • Eliminates the light stimulus for weed seed germination over most of the planting bed.
  • Blocks emergence of most weeds.
  • Conserves soil moisture present at the time of installation or provided by drip lines under the mulch.
  • Minimizes nutrient leaching by shedding excessive rainfall.
  • Raises soil temperatures by a few degrees, thereby promoting early-season crop growth and maturation.
  • Helps keep the edible portion of vegetable crops clean, especially pumpkin and other fruiting vegetables.

However, plastic mulch also:

  • Is manufactured from petroleum, a non-renewable resource.
  • Does not provide organic matter to feed the soil.
  • Does not provide as good a habitat for ground beetles, earthworms, and other beneficials as organic mulches.
  • Does not breathe, excludes rainfall, and requires drip irrigation to ensure adequate moisture.
  • May require manual removal of weeds emerging through planting holes.
  • Can be penetrated by nutsedges and a few other weeds with sharp, tough growing points.
  • Requires cultivation or other measures to control weeds in alleys.
  • Can channel runoff and worsen between-bed erosion during heavy rains on sloping fields.
  • Must be picked up and disposed of at the end of the season.
  • Generates large volumes of plastic waste (200–300 lb/ac).

Despite these drawbacks, many farmers use plastic mulch because it is well suited to mechanized, medium- to large-scale production. The yield increases and labor savings from using black plastic often pay for the purchase price and pickup costs several times over. Ensuring a tight fit of plastic film to the soil surface, through precise bed preparation and mulch application, improves weed control and enhances soil warming (Fig. 2). Drip irrigation is important, as laying plastic without practical means to irrigate the crop can result in moisture deficit and yield losses (Schonbeck and Evanylo, 1998).

Installing and planting plastic mulch
Figure 2. (a) Black plastic mulch and drip lines are laid mechanically on precisely formed raised beds to give a tight fit. (b) A single pass operation makes planting holes, sets tomato starts, and delivers liquid organic starter fertilizer. Figure credits: (a) Jim Shrefler, Oklahoma State University; (b) Becky Crouse, Marketing Manager, Potomac Vegetable Farms, Purcellville, VA.

Black plastic mulch does not eliminate all weeds. Light reaching the soil surface through planting holes and in uncovered alleys between mulched beds allows weeds to emerge and establish (Fig. 3); thus, additional measures are needed to manage weeds in these areas. Weeds in plant holes can compete severely with the crop if they are not removed early (Fig. 4a), and vining species like morning glories (Ipomoea spp.) and bindweeds (Convovulus arvensis and Calystegia sepium) grow toward the light of planting holes and climb the crop plant (Fig. 4b). Nutsedges (Cyperus spp.) and a few other weeds with sharp growing points can puncture and emerge through the film. Black plastic can accelerate the growth and spread of purple nutsedge (C. rotundus) by warming the soil (Webster, 2005).

Weeds in plastic mulch
Figure 3. Weeds have begun to grow in the unmulched alleys between plastic-mulched beds. In addition, some weeds have emerged through planting holes, and a few have punctured the film itself. Photo credit: Becky Crouse, Marketing Manager, Potomac Vegetable Farms, Purcellville, VA.

Weed growth through planting holes in plastic mulch
Figure 4. (a) Pigweed, emerged through the planting hole, is now too large to remove by pulling, and is competing significantly with the eggplant crop. (b) Ivyleaf morning glory has emerged through the planting hole, and has begun to climb the potato crop. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Weed growth in alleys and along the edges of the mulch can be a challenge to manage. Tactics include cultivating, flaming, mowing, organic mulching, or cover cropping. Cultivation requires care to avoid cutting the mulch or pulling it loose above or below ground. Many growers apply straw, hay, or other organic mulch to the alleys and overlap it onto the plastic (Fig. 5), either at the time of planting, or after one or more cultivations. The organic mulch suppresses alley weeds, conserves soil moisture, and provides organic matter that can later be incorporated into the soil. Cover crops such as buckwheat or clover can be planted in alleys, either when the plastic is laid, or after one or more alley cultivations. At least one grower has had success with planting a ryegrass cover crop in alleys immediately after the mulch is installed, thereby eliminating the need for cultivation (Josh Hardin, farmer in Arkansas, pers. commun). Alley cover crops can be maintained by mowing to minimize shading of the vegetable and facilitate harvest (Fig. 6).

Using organic mulches with plastic mulches
Figure 5. A thick layer of hay applied to alleys between plastic-mulched rows of eggplant has suppressed most alley weeds. With the onset of hot summer weather, some hay was moved onto the plastic to prevent excessive soil heating. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Using cover crops with plastic mulch
Figure 6. A cover crop of rye and clover, planted when the plastic mulch was laid and maintained by mowing, adds organic matter, protects the soil, and facilitates access to the crop for disabled adults who work at Red Wiggler Farm in Maryland. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

For best results with black plastic mulch:

  • Avoid using plastic in fields heavily infested with nutsedge or vining weeds.
  • Prioritize early plantings and heat-loving crops for use of black plastic.
  • Add organic nutrient sources and other amendments to the bed before applying plastic.
  • Install in-row drip irrigation tape under the film in such a way that the tape will not be damaged during planting operations (burial to the side of the row is often done).
  • Ensure a tight fit of plastic film to soil surface to improve weed control and soil warming.
  • Manage alley weeds through timely cultivation, mowing, mulching, or cover crops.
  • If soil temperatures under plastic exceed optimal ranges for the crop, spread organic mulch over plastic.
Clear, Translucent, and Colored Plastic Films

Clear, translucent, and infrared-transmitting (IRT) plastic films that allow solar radiation to reach the soil will warm the soil more effectively than does black plastic. However, clear colorless plastic may allow such vigorous weed growth that the mass of vegetation bulges and eventually tears the plastic. During very hot sunny weather, a tight fitting clear plastic film can heat the soil sufficiently to kill rhizomes and other vegetative weed propagules, some weed seeds, and most plant pathogens in the uppermost several inches of the soil profile. This process is called soil solarization, and is a valuable tool for preparing small areas for planting certain high-value crops. Vegetables are not normally planted into clear plastic, because the heat buildup under this mulch can damage the crop during hot weather, and accelerate weed growth during cooler conditions.

Translucent green, brown, olive, and IRT (infrared-transmitting) plastic films (Fig. 7) have been developed, which combine greater soil warming (compared to black plastic), with fair weed suppression. Because they absorb the red and blue light wavelengths used by all plants in photosynthesis, and transmit mostly infrared (heat) wavelengths and some green light, these materials reduce weed germination, emergence, and growth compared to clear film or bare soil. Translucent films do not control weeds as effectively as black plastic, and should be used on crops for which soil warming is critical, and in fields with light-to-moderate weed populations.

IRT film mulch
Figure 7. A translucent, infrared-transmitting (IRT) film mulch promotes greater soil warming for this pepper crop without transmitting enough visible light to support rapid weed growth. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Film mulches with white or reflective surfaces lower soil temperature compared to uncovered soil. This can benefit crop growth and competitiveness against weeds during the hottest summer months, when soil temperature under black mulch would exceed the optimum range for the crop. For example, tomato prefers moderately warm (70–85 °F) root zone temperatures and can be stressed by higher temperatures (Tindall et al., 1990); thus, farmers in warm climates often use a white mulch for later tomato plantings, or apply whitewash to black mulch when hot weather sets in after the crop is planted.

Research has shown that the quality of light reflected by mulch can affect crop production (Orzolek and Lamont, 2000). Reflective (silver colored, aluminum-coated) films (Fig. 8) disorient and repel aphids, whiteflies, and some other pests. Red plastic has been reported to enhance tomato and strawberry yields by 12–20%, and dark blue plastic to improve cucurbit yields (Orzolek and Lamont, 2000). In South Carolina, red mulch was found to stimulate tomato fruit growth (Kasperbauer and Hunt, 1998), and reduce pest nematode attack on the crop (Adams, 1997). However, studies in Iowa showed no difference in tomato yield in red, IRT, and black mulches (Taber and Smith, 2000). Lack of consistency among research results suggests that the benefits of different colored mulches may vary with local conditions (climate, insect pest populations, etc.); thus, farmers may be well advised to test specialized colored mulches on a small scale before investing in application on a larger area.

Reflective plastic mulches
Figure 8. Reflective plastic mulches. (a) A reflective, aluminum-coated, opaque plastic film suppresses weeds while also repelling aphids and other pests in this cucumber crop. (b) Reflective mulch keeps the soil cooler for pepper, eggplant, and tomato during hot summer weather than does black plastic (right background). Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

White-on-black and red-on-black film mulches have been developed to combine the weed suppression of a completely opaque mulch with the yield and pest management benefits of colored mulch.

Weed Barrier and Landscape Fabric

Woven black polypropylene mulches, marketed as weed barrier, weed fabric, or landscape fabric, provide a durable and effective barrier to weed growth, and are especially popular for perennial plantings. Used alone, they promote soil warming similar to black plastic film; if cooler soil conditions are desired, the mulch can be covered with a layer of chipped brush or other organic mulch. Spun fabric mulches are readily penetrated by quackgrass (Elytrigia repens) and other rhizomatous perennials, and are not recommended (Mohler and DiTommaso, unpublished). Often used in ornamental plantings, woven fabric mulches have found increasing use in commercial horticultural food crop production, especially berries and other high-value perennial crops, because they are:

  • Permeable to air, water, and nutrients.
  • Opaque and durable, giving effective weed suppression.
  • Long lasting, typically 8–12 years.

Disadvantages are that the materials are relatively heavy (2.4–4.1 ounces per square yard), and expensive, although the cost is amortized over many years' use. USDA Organic Standards require that the mulch be removed from the field at the end of its useful lifespan.

Some growers use woven mulches for vegetable crops (Fig. 9), reusing the same material for multiple crops and seasons. For example, Hiu Newcomb of Potomac Vegetable Farms in Vienna, VA has found this material economical for small-scale cucumber production, and picks it up for reuse as soon as harvest is complete (pers. commun.).

Landscape fabric
Figure 9. A heavy-duty landscape fabric has greatly reduced weed pressure in potato. Covering the black fabric with straw or other light colored organic material at the onset of hot weather is recommended to maintain soil temperatures favorable for potato tuber development. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Biodegradable Plastic Mulch?

For years, researchers have tried to develop degradable plastic mulches that would eliminate the need to gather and dispose of spent, dirty plastic at the end of the season, which can entail significant costs for medium-to-large scale operations. Many growers rejected the early versions of photodegradable and biodegradable mulches, which left fragments of undecomposed mulch in the soil and thereby created a litter problem. More recently, plant-starch-based biodegradable black plastic films have been developed. These perform as well as standard black polyethylene film, and appear to decompose completely (Rangarajan and Leonard, 2007; Ngouajio et al., 2008). However, as of the fall of 2011, the USDA National Organic Program (NOP) has not yet approved these products, which contain synthetic ingredients, residues of which could remain in the soil after the mulch has broken down.

Paper mulch

Farmers and researchers have experimented with paper mulches as an alternative to plastic, with mixed results. In one study, paper mulches slightly lowered soil temperatures, and tended to tear along soil-anchored edges (Fig. 10), resulting in slightly lower yields and less effective weed control than with black plastic (Schonbeck, 1998). Since then, several commercial biodegradable paper mulch products have been approved by NOP (Organic Materials Review Institute), and have performed fairly well in field trials (Miles et al., 2007). Farmers may need to add more soil to the edges at midseason to keep the paper mulch anchored.

Paper mulch
Figure 10. In this field trial, a black recycled paper mulch product began to break down before the end of the season, allowing some weeds to emerge in the crop. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Whereas paper alone may not always give adequate weed control, a single layer of mulch paper or several layers of newsprint under a layer of hay or other organic mulch can substantially enhance weed suppression (Fig. 11). Paper + 5 tons per acre of hay suppressed aggressive weeds like Bermuda grass (Cynodon dactylon) and common cocklebur (Xanthium strumarium) as effectively as 10 tons per acre of hay without paper (Schonbeck, 1996 and 1998). NOP allows the use of newspaper and cardboard for mulch provided that they are non-glossy and free of colored inks.


Figure 11. In this tomato planting in Floyd, VA (Appalachian region), a combination of several layers of newspaper plus hay mulch adequately controlled weeds, including the rhizomatous perennial quackgrass (Elymus repens), throughout the season. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

References Cited
  • Adams, S. 1997. Red plastic mulch thwarts nematodes. News and Events. United States Department of Agriculture–Agricultural Research Service. (Available online at: http://www.ars.usda.gov/is/pr/1997/971003.htm) (verified 10 Sept 2012).
  • Kasperbauer, M. J., and P. G. Hunt. 1998. Far-red light affects photosynthate allocation and yield of tomato over red mulch. Crop Science 38: 970–974. (Available online at: https://www.crops.org/publications/cs/abstracts/38/4/CS0380040970) (verified 10 Sept 2012).
  • Miles, C., E. Klingler, L. Nelson, T. Smith, and C. Cross. 2007. Alternatives to plastic mulch in vegetable production systems. Research Report, Washington State University. (Available online at: http://vegetables.wsu.edu/MulchReport07.pdf) (verified 10 Sept 2012).
  • Mohler, C. L., and A. DiTommaso. Unpublished. Manage weeds on your farm: A guide to ecological strategies. Version 5.1 (4 Dec 2008).
  • Ngouajio, M., R. Auras, R. T. Fernandez, M. Rubino, J. W. Counts, Jr., and T. Kijchavengkul. 2008. Field performance of aliphatic–aromatic copolyester biodegradable mulch films in a fresh market tomato production system. HortTechnology 18: 605–601. (Available online at: http://horttech.ashspublications.org/content/18/4/605) (verified 10 Sept 2012).
  • Orzolek, M. D., and W. J. Lamont, Jr. 2000. Summary and recommendations for the use of mulch color in vegetable production. (Available online at: http://extension.psu.edu/plasticulture/technologies/plastic-mulches/summary-and-recommendations-for-the-use-of-mulch-color-in-vegetable-production) (verified 10 Sept 2012).
  • Rangarajan, A., and B. Leonard. 2007. Biodegradable mulches: How well do they work? (Available online at: www.newenglandvfc.org/pdf_proceedings/biomulches.pdf) (verified 10 Sept 2012).
  • Schonbeck, M. W. 1996. Mulching for weed control in annual vegetable crops. Virginia Association for Biological Farming Information Sheet.
  • Schonbeck, M. W. 1998. Weed suppression and labor costs associated with organic, plastic, and paper mulches in small-scale vegetable production. Journal of Sustainable Agriculture. 13: 13–33. (Available online at: http://dx.doi.org/10.1300/J064v13n02_04) (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).
  • Taber, H. G., and B. C. Smith. 2000. Effect of red plastic mulch on early tomato production. (Available online at: www.public.iastate.edu/~taber/Extension/Progress Rpt 00/redmulch.pdf) (verified 10 Sept 2012).
  • 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).
  • Webster, T. M. 2005. Patch expansion of purple nutsedge (Cyperus rotundus) and yellow nutsedge (Cyperus esculentus) 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).

 

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 4872

Spiny Amaranth (Amaranthus spinosus)

mer, 2012/09/12 - 16:57

eOrganic author:

Dr. Mark Schonbeck, Virginia Association for Biological Farming

Introduction

Spiny amaranth, sometimes called spiny pigweed, is a troublesome weed of vegetables, row crops, and pasture in warm climates. Native to the lowland tropics in the Americas, spiny amaranth has spread through tropical and subtropical latitudes around the world (Holm et al., 1991), has become a major weed of rice in the Philippines (Chauhan and Johnson, 2009), and is moving into temperate regions in the United States. Its widespread distribution and its sharp spines, which deter grazing and interfere with manual weeding and harvest, have earned spiny amaranth designation as the world's 15th worst agricultural weed (Holm et al., 1991).

Spiny amaranth is an erect, often bushy, much-branched summer annual, growing to heights of 2–5 feet (Fig. 1). Stems and leaves are smooth and hairless, sometimes shiny in appearance. Each leaf node along the stem bears a pair of rigid, sharp spines ~0.5 inch long (Fig. 2).

Mature piny amaranth
Figure 1. A large, mature specimen of spiny amaranth, showing its bushy habit of growth. Photo credit: John D. Byrd, Mississippi State University, Bugwood.org.

Spiny amaranth spines
Figure 2. Spiny amaranth shoot in early flowering, illustrating sharp spines at each leaf node. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Leaf blades are egg-shaped to diamond-shaped, with the broader end closest to the stem, and 1–4 inches long by 0.5–2.5 inches wide. The petiole is approximately as long as the blade. Leaves may be variegated with a v-shaped watermark or area of lighter color (Fig. 3), although this is not a definitive characteristic of this species, since some other amaranths can show a similar watermark.

Spiny amaranth seedlings
Figure 3. (a) A flush of spiny amaranth seedlings emerging in alley between crop beds. Note v-shaped variegation on leaves of older seedlings at upper left. (b) Young spiny amaranth with pronounced leaf variegation, in Maui, Hawaii. Figure credits: a Mark Schonbeck, Virginia Association for Biological Farming. (b) Forest and Kim Starr, U.S. Geological Survey, Bugwood.org.

Like other pigweeds, spiny amaranth develops a strong taproot with a network of fibrous feeder roots. The taproot may or may not be distinctly reddish in color.

Male and female flowers are borne on different regions of the same plant—linear or branched terminal spikes with mostly male flowers, and globular axillary clusters of mostly female flowers (Bryson and DeFelice, 2009) (Fig. 4).

Spiny amaranth flowers
Figure 4. Spiny amaranth inflorescences include mostly male terminal spikes, and small globular clusters of mostly female flowers at leaf nodes. Photo credit: Mark Schonbeck, Virginia Association for Biological Farming.

Biology

Spiny amaranth thrives in rich, warm, loamy soils with high organic matter and sufficient nitrogen (N), and can produce up to 235 thousand seeds per plant (Holm et al., 1991). In cooler regions, spiny amaranth does not grow as rapidly as other pigweeds. In a field study in Missouri, spiny amaranth reached 4.5 feet and produced 114 thousand seeds per plant within 14 weeks, compared to 5–7 feet and 250–292 thousand seeds for several other weedy Amaranthus species (Sellers et al., 2003).

Flowering in spiny amaranth has been described as day-neutral (Holm et al., 1991), unlike other pigweeds, which flower in response to shortening days in late summer. Although the weed is most troublesome in warm or tropical regions, different ecotypes are known to exist, some of which flourish in temperate agricultural ecosystems.

The seeds of spiny amaranth are very small (~0.03 inch diameter), which is smaller than the seeds of some other pigweeds (Uva et al., 1997). Seeds may be carried by wind, water, or animal dung (Holm et al., 1991). Many farmers in Virginia and neighboring states report that spiny amaranth has arrived on their farms in livestock manure (personal communication).

Spiny amaranth seeds germinate mostly from the soil surface, and burial in the soil to a depth of just 0.25 inch can reduce germination considerably (Chauhan and Johnson, 2009). The seeds germinate in warm conditions (68–95 °F), and they respond strongly to diurnal temperature fluctuations and to light (Steckel et al, 2004; Chauhan and Johnson, 2009). Germination and emergence may take a few days longer for spiny amaranth than for some other amaranths (Sellers et al, 2003; Steckel et al., 2004).

Impact on Crops

Spiny amaranth appears to be a moderately strong competitor in U.S. croplands. It is most likely to cause yield losses in warmer parts of the South, and in shorter-statured crops that cannot shade out the weed. Because of its stiff, sharp spines, which can draw blood, spiny amaranth is a major nuisance in vegetables and other crops that are tended, weeded, and harvested manually. Grazing livestock are deterred by the spines, and the weed can become toxic when growing in soils high in available N. Unlike other pigweeds, which are highly palatable to livestock, spiny amaranth can become a significant weed in pasture or rangeland (Chauhan and Johnson, 2009) (Fig. 5).

Spiny amaranth in pasture
Figure 5. (a) Spiny amaranth growing in pasture in northwest Arkansas. (b) Spiny amaranth infestation in pasture. Figure credits: (a) Mark Schonbeck, Virginia Association for Biological Farming; (b) Chris Evans, River to River CWMA, Bugwood.org.

In field and greenhouse studies on an organic (muck) soil of the Florida Everglades, low levels of available phosphorus limited the growth and yield of lettuce, but not of spiny amaranth (Santos et al., 1997; Shrefler et al, 1994). P fertilization improved the competitiveness of lettuce against the weed, and band application of the fertilizer reduced spiny amaranth impact on yield.

Management Implications for Organic Production

Organic vegetable and livestock producers who do not already have spiny amaranth on their farms will want to avoid importing this nuisance. Manure from off-farm sources should be hot composted at 140–160 °F (ideally 160 °F) for two weeks with several turnings to kill weed seeds, before field application.

If spiny amaranth is already present, or field scouting reveals that it has arrived, utilize the management strategies described for pigweeds. Because of its very small seed and seedling size, shallow depth of emergence, and germination response to temperature fluctuations, spiny amaranth may be especially susceptible to the following tactics:

  • Shallow and early cultivation when the weed is small and the taproot is easily severed or disrupted
  • Burial of within-row weeds by hilling-up
  • Organic mulch (physical hindrance, dampening thermal germination stimuli)
  • One-time moldboard plowing to bury the seed bank
References Cited
  • Bryson, C. T., and M. S. DeFelice. 2009. Weeds of the South. University of Georgia Press, Athens, GA.
  • Chauhan, B. S., and D. E. Johnson. 2009. Germination ecology of spiny (Amaranthus spinosus) and slender amaranth (A. viridis): Troublesome weeds of direct-seeded rice. Weed Science 57: 379–385. (Available online at: http://dx.doi.org/10.1614/WS-08-179.1) (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.
  • 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.
  • 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).
  • 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).
  • Uva, R. H., J. C. Neal, and J. M. DiTomaso, 1997. Weeds of the Northeast. Cornell University Press, Ithaca, NY.

 

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 5125

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
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  • 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).
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  • 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).
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  • 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).
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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
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Organic Vegetable Production Systems, Control Practices in Organic Weed Management

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Organic Vegetable Production Systems, Production of Specific Vegetable Crops

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Control Practices in Organic Weed Management

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