|
http://www.ag.ohio-state.edu/~corn/ |
|
September 17 to September 23,
2001
C.O.R.N. 2001-31
In This Issue:
A) Managing Winter Annual Weeds with Fall Herbicide Applications
B) Yield Monitor Calibration Time
C) Commonly Used Methods for Detecting GMOs in Grain Crops
D) Receive a Free Gift at Farm Science Review
Populations of winter annual weeds seem to have been at an all time high over the past several years. Weeds such as common chickweed, henbit, purple deadnettle, and marestail (horseweed) have increased to the point that they require changes in herbicide management in some fields. Several factors may have caused the increase in winter annual weeds. Most winter annuals emerge in the fall, and the warm weather in late fall during the past several years has resulted in higher populations. No-tillage tends to promote winter annual populations, since there is no tillage in fall to disrupt their emergence. However, winter annuals can emerge after an early fall tillage, and have been a problem in tilled as well as no-till fields. Early soybean harvest in 1999 and 2000 allowed earlier than typical fall tillage in some fields, providing a window after tillage for winter annual emergence. Another factor may be the switch from preplant/preemergence herbicide programs (Squadron, Canopy etc) to Roundup Ready and other postemergence programs, since we have observed winter annual weed problems showing up more often following postemergence programs. If so, this may indicate that the preplant herbicides are either: 1) preventing seed production by these weeds in the spring, or 2) persisting into the fall at rates that are high enough to reduce winter annual emergence. Continued problems with winter annuals may warrant reconsideration of the utility of total postemergence programs.
The Life Cycle of Winter Annual Weeds
Chickweed, deadnettle, and most other winter annual weeds emerge primarily in
the late summer or fall, although some emergence can occur in the spring. Marestail
and common chickweed can follow a winter annual or summer annual life cycle,
and has the potential to emerge over most of the year depending upon environmental
conditions. Winter annuals survive the winter with little or no further growth,
resume growth in late-winter or early spring, and flower and go to seed in late-spring
or early summer. Consequently, the major negative effect of most winter annuals
occurs at the time of crop establishment. Marestail does not flower until late
summer, and directly competes with crop growth along with foxtail, ragweeds,
and other summer annuals. We have occasionally observed completion of the winter
annual life cycle within a shorter period of time. For example, when chickweed
gets an early start in late summer, it may set seed by late fall or early winter.
One characteristic of winter annuals that allows populations to increase over
a short time period is the lack of after-ripening needed for seed viability.
While seed from many weed species requires a year or more of after-ripening
in the soil in order to germinate, seed from winter annuals is typically ready
to germinate as it leaves the plant.
Problems Caused by Dense Populations of Winter Annual Weeds
Purple Deadnettle and Soybean Cyst Nematode
OSU research has shown that some winter annuals can serve as hosts for soybean
cyst nematode. In Ohio, the primary potential host appears to be purple deadnettle.
OSU research indicates a possibility that deadnettle emerging in late summer
could serve as an alternate host for cyst nematode when the soybeans senesce,
allowing completion of another nematode life cycle and increasing their populations.
We are therefore suggesting that in fields where cyst nematode is a known problem,
deadnettle should be controlled by late September or within 30 days of deadnettle
emergence to interrupt the nematode life cycle. Have fields tested for the amount
of soybean cyst nematode if you are unsure whether it is a problem.
Strategies for Fields Where Winter Annuals are a Problem
Since the major problems with chickweed and deadnettle often are the slow soil
drying and interference with crop planting and tillage, the primary goal of
winter annual management should be to allow maximum time for weed death and
dessication. This may be accomplished with fall or early spring herbicide applications
or tillage. Applications too soon before planting may not kill plants rapidly
enough, especially during periods of cold weather. Control can be achieved with
tillage or herbicides, although herbicides will be the method of control in
no-till fields. Some advantages and disadvantages of the various management
strategies:
Are All Fields Candidates for Fall Herbicide Programs?
Winter annual populations vary greatly among no-till fields, and some fields
have few winter annuals at this point. We do not see a great advantage to fall
applications in these fields. Any type of program that has been consistently
effective in prior years, including application of herbicides at planting for
burndown, can still be used. Since there appears to be a general trend for increasing
winter annual populations, however, these fields should be scouted each fall
to ensure that populations are not increasing. Preventing seed production in
low populations of winter annuals will minimize the risk of future problems.
Overview of OSU Research on Winter Annuals
OSU Weed Scientists conducted field research in the fall and spring of 1999/2000
and 2000/2001 to determine the effectiveness of various herbicide treatments
for control of common chickweed and purple deadnettle. We had field studies
at three locations each year east of Chillicothe, Amanda, and South Charleston.
Our research was much more extensive in 2000/2001, with regard to the number
of herbicides and rates included. Fall herbicide treatments were applied in
mid- to late-November both years, and spring treatments were applied in late
March. Results of these studies are summarized below and tables showing all
of the data are available at the OSU Weed Science website
http://www.oardc.ohio-state.edu/weedworkshop/
We included 2,4-D ester (1 pint/A) with most treatments. Treatments containing glyphosate were applied with ammonium sulfate. All other treatments were applied with crop oil concentrate. Overall, fall treatments provided much more consistent control of chickweed and deadnettle than spring treatments. Among soybean herbicides, few treatments provided more than 80% control of both weeds in March, but a number provided better than 90% control when applied in November. Most of the chickweed and deadnettle (95% or greater) emerged prior to November treatments, but we did observe a few deadnettle emerging in the spring where treatments without residual activity were applied.
Fall herbicide treatments for soybeans providing an average of at least 90%
control of chickweed and deadnettle in OSU research:
Fall herbicide treatments for soybeans providing an average of 80 to 90% control
of chickweed and deadnettle in OSU research:
Spring herbicide treatments for soybeans providing an average of 80 to 92%
control of chickweed and deadnettle in OSU research:
Fall herbicide treatments for corn providing an average of at least 90% control
of chickweed and deadnettle in OSU research (Note: simazine + 2,4-D ester is
effective on most winter annuals but is weak on deadnettle):
Spring herbicide treatments for corn providing an average of at least 90% control
of chickweed and deadnettle in OSU research (glyphosate and Gramoxone were not
included in corn trial, but we assume results would be similar to soybean study):
Some Suggestions for the use of 2,4-D Ester
We included 2,4-D ester in most treatments because it is economical and helps
control dandelion, marestail, and mustards. 2,4-D is generally not effective
for control of chickweed or deadnettle, but can have some activity on these
weeds. Where dandelion is target weed, use a 2,4-D rate of 1 quart/A unless
it is combined with Canopy XL or glyphosate, in which case a rate of 1 pint/A
may be adequate.
Fall Herbicide Options for Dandelion
Dandelion control in the spring has been variable and extremely slow with many
herbicide treatments, especially when the dandelions are well established. Fall
is an excellent time to apply herbicides for dandelion control. We suggest applying
by early November when control of dandelion is a goal. Herbicide options include:
Fall Herbicide Options for Marestail (Horseweed)
Marestail can emerge throughout much of the year, but the majority emerges in
the fall. Fall herbicide options include:
Suggestions on Adjuvants
Application Timing
Data from OSU field studies shows that application in the fall from about mid-October
to early December can provide effective control of winter annuals. Herbicides
should be applied by early November for control of dandelion and Canada thistle.
Where winter annual populations are dense in the spring, applications should
be made by late March to allow time for weed dessication. Marestail becomes
more difficult to control as it increases in size, so fall or early spring applications
are more effective than applications at planting. Glyphosate is more inconsistent
in the spring for control of marestail.
Need for a Burndown Application at Planting
Use of herbicides in the fall does not necessarily eliminate the need for an
application of burndown herbicides at planting the following spring. This will
be affected by the nature of the weed populations in the field, date of planting,
and type of herbicides used in the fall. While a fall application may control
most of the winter annual weeds, some summer annual weeds start to emerge in
early spring. Early-emerging weeds can include ragweeds, smartweed, atriplex,
and lambsquarters. When herbicides with residual activity (Backdraft, Canopy
XL, Sencor, Python) are applied in the fall, they should provide at least some
control of these weeds into the following spring. Fall treatments without residual
(glyphosate, paraquat, 2,4-D) provide no control of spring-emerging weeds. Based
on our experience in conventional and Roundup Ready soybeans, we suggest the
following:
http://precisionag.osu.edu/library/ymonitor.html
Prior To Calibration
Using Scales and Calibrating Loads
For more tips about yield monitor calibration contact your yield monitor manufacturer or local dealer. Additional resources, information and news about precision agriculture can be found at The Ohio State University's Precision Agriculture Web Site http://precisionag.osu.edu
As market restrictions for various transgenic (genetically modified organism or GMO) crops (e.g. Bt-corn, Roundup Ready soybeans and corn) continue, there is increasing interest among growers in determining the presence of GMOs in crops. Growers producing non-GMO grains for specialty markets need to verify that there is no GMO contamination or that contamination levels meet tolerance levels established by an end user. The default standard for certification as GMO free has been taken to be zero in many cases, although experience shows that meeting such a standard will be difficult. There have been proposals for setting maximum allowable levels in the range of 1 to 3%, and it's likely that some tolerance level above zero will be accepted in the future. Japan recently established new legislation that sets a zero tolerance for seed and food imports containing unapproved biotech material, e.g. StarLink corn (containing the Cry9C Bt transgene). The Japanese legislation will allow food products containing less than 5 % of approved biotech crops like corn and soybeans to be labeled as non-GMOs. The European Union (EU) has recently proposed rules on the labeling and traceabililty of foods containing GMOs. According to these new rules, accidental traces of GMOs that have been cleared by the EU's scientific advisers, even if they have not received final official approval, will be allowed in food and feed up to a maximum of 1% without being subject to labeling requirements. Tolerances similar to those of Japan and the EU are being considered by other countries importing U.S. grains.
There are also other circumstances when GMO testing may be useful in crop production, such as when troubleshooting crop disorders during the growing season. If an allegedly herbicide resistant corn or soybean field exhibits extensive injury following herbicide application, the grower may want to confirm that plants in the field are actually herbicide resistant. Similarly a grower, if uncertain, may need to determine what fields, or what parts of fields, he planted to GMO crops.
There are several commonly used GMO testing protocols, including biological tests, as well as ELISA and PCR tests, for herbicide and insect tolerance. Growers and end users should consider the advantages and disadvantages of the various testing methods before harvest. Exporters should probably resign themselves to the most rigorous testing protocol to anticipate the additional scrutiny their products will receive overseas. Some major end-users, i.e. large food processors, are currently using a combination of tests for identity-preserved (IP) grains.
Herbicide bioassays are used to detect GMO herbicide resistant traits in Roundup Ready and Liberty Link soybeans and corn. The tests involve placing seeds in a germination media moistened with a diluted solution containing the herbicide or spraying the herbicide on seedlings. Seeds that test positive for the presence of the herbicide tolerant GMO trait will germinate and develop normally, whereas those that die or do not develop normally will be GMO-free. This procedure is widely used by seed and grain companies exporting soybeans. Advantages of the bioassay method: it's relatively inexpensive ($20 -$30), user friendly, and produces straightforward results. Disadvantages of the test: it takes up to a week to complete, its use is limited to the Roundup Ready and Liberty Link herbicide resistant GMO crops, and the seeds need to germinate for the test to work. Herbicide bioassays can also be used to detect herbicide resistant traits in non-GMO corns such as Clearfield® hybrids which are tolerant to imidazolinone herbicides (Pursuit, Scepter, Lightning).
ELISA (enzyme-linked immunosorbent assay) tests for the presence of the specific protein that the genetically modified DNA produces in the plant. ELISA procedures use antibodies that react with specific proteins produced by the GMO. There are different versions of the ELISA method used for GMO detection. One version uses lateral flow strips and delivers results in 2 to 5 minutes. This "strip test" technology is similar to that used in home pregnancy tests. Strip tests are commonly used at grain elevators where a rapid assessment to determine the presence or absence of GMOs is needed. These tests are referred to as the "dipstick" procedure by some companies marketing this ELISA technology for seed testing. Another version of the ELISA test, the "plate test", provides some indication of the quantity (percentage) of the tested sample that is the GMO in question. Intensity of color indicates the amount of the protein present. The plate test can take 2 to 4 hours and is more laborious, and costly than the strip test. Advantages of the ELISA strip tests are speed, relative ease of use, and low cost. The major disadvantage of the strip test is that it cannot quantify how much GMO is present. ELISA tests have limited application for testing GMOs in processed foods because heat processes denature the proteins, thereby making detection of proteins difficult.
The PCR (polymerase chain reaction) method is more sensitive than the ELISA method and tests for the presence of the specific DNA sequence of the gene itself. The major advantage of PCR tests is sensitivity, i.e. detection of GMOs at very low levels. PCR is the only one of these methods that can effectively detect GMOs in processed foods. Major disadvantages of the PCR protocol include length of time needed (2-3 days), and cost ($75-$300 per sample). PCR tests also require more sophisticated equipment and greater expertise. While more sensitive to GMOs, PCRs in some cases tend to show false positives. PCR procedures were originally developed as research tools for analyzing genes and assisting in the movement of genes among organisms. Given the expense, time, and expertise required, PCR testing has limited potential in the field or at grain elevators.
As the number of GMO traits increases (e.g. GMO corns with resistance to Roundup, European corn borer, and western corn rootworm), it will become more costly to monitor the presence of GMOs in crops, since each different gene requires a separate test. However, if the demand for non-GMO crops increases, it's possible that tests for different genes may be combined on the same ELISA test strip.
Although some of these GMO testing procedures such as the ELISA strip tests can be used in the field and elevators, the other procedures for detecting GMOs require more sophisticated training and equipment to be used effectively. The following is list of some of the laboratories that offer GMO testing of grain crops for a fee. The websites for these labs provide an overview of the specific GMO testing procedures they conduct. The following are some commercial laboratories testing for GMOs in crops (as of 7-10-01):
Central Hanse Analytical Lab LLC
101 Wordland Hwy
Belk Chasse, LA 70037
504-393-5290; fax 504-393-5270
www.rmgcal.com/cal
Biogenetic Services Inc.
801 32nd Ave
Brookings, SD 57006
605-697-8500; fax 605-697-8507
www.biogeneticservices.com
California Seed & Plant Lab, Inc.
7877 Pleasant Grove Rd
Elverta, CA 95626
916-655-1581; fax 916-655-1582
www.calspl.com
Genetic ID
1760 Observatory Dr
Fairfield, IA 52556-9030
888-229-2011; fax 641-472-9198
www.genetic-id.com
Mid-West Seed Services
236 32nd Ave
Brookings, SD 57006
605-692-7611; fax 605-692-7617
www.mwseed.com
Ohio Seed Improvement Association
6150 Avery Rd, Box 477
Dublin, OH 43017-0477
614-889-1136; fax 614-889-8979
www.ohseed.org
AGDIA
30380 Country Road 6
Elkhart, IN 46514
219-264-2014; fax 219-264-2153
www.agdia.com
Biodiagnostics Inc.
507 Highland Dr
River Falls, WI 54022
715-426-0246; fax 715-426-0251
Indiana Crop Improvement
770 Stockwell Rd
Lafayette, IN 47909
765-523-2535; fax 765-523-2536
www.indianacrop.org
Illinois Crop Improvement
3105 Research Rd, PO Box 9013
Champaign, IL 61826-9013
217-359-4053; fax 217-359-4075
www.ilcrop.com
OTHER REFERENCES RELATING TO GMO TESTING
Nafziger, E. 2000. GMOs: Q & A. University of Illinois. http://www.ag.uiuc.edu/~stratsoy/expert/gmo.html
The Non-GMO Source. 2001. An Overview of GMO Testing Methods. 1:4
Biotechnology Issues on the Web.2001 Purdue University. http://www.agry.purdue.edu/ext/corn/cafe/biotech.html
The first 25 visitors each day to the Ohio State University Agronomic Crops Team tent at the Farm Science Review will receive a copy of Bulletin 827, Corn, Soybean, Wheat and Alfalfa Field Guide as a free gift. Stop by the tent and mention that you read about this offer in the C.O.R.N. Newsletter to receive your Field Guide. The Agonomic Crops Team display tent is located on Friday Avenue, across from the Bailey and Firebaugh Buildings and next to the Ohio Farmer magazine exhibit.
While at the Crops Team exhibit, take a few minutes to try a weed identification quiz, and look at displays on the soybean aphid, soybean cyst nematode, sudden death syndrome in soybeans, and weed resistance in Ohio. Displays from the OSU Extension Precision Agriculture Team will also be at the same location.
Readers can subscribe electronically to this newsletter by sending an e-mail message to: corn-out-on@postoffice.ag.ohio-state.edu. A successful subscription message will receive by an automatic reply from the listserv. Contact your local Ohio State University Extension Office or e-mail labarge.1@osu.edu if you have problems subscribing.
Past versions of C.O.R.N. can be found on the World Wide Web at: http:/www.ag.ohio-state.edu/~corn/archive/
C.O.R.N. is a summary of crop observations, related information, and appropriate recommendations for Ohio Crop Producers and Industry. C.O.R.N. is produced by the Ohio State University Extension Agronomy Team, State Specialists at The Ohio State University and Ohio Agricultural Research and Development Center. C.O.R.N. Questions are directed to State Specialists, Extension Associates, and Agents associated with Ohio State University Extension and the Ohio Agricultural Research and Development Center at The Ohio State University.
State Specialists: Pat Lipps, Anne Dorrance and Dennis Mills (Plant Pathology), Peter Thomison (Corn Production), Ron Hammond and Bruce Eisley (IPM), Mark Loux and Jeff Stachler (Weed Science) Extension Agents: Barry Ward (Champaign), Howard Siegrist (Licking), Ray Wells (Ross), Roger Bender (Shelby), Gary Wilson (Hancock) and Andy Kleinschmidt (Van Wert)Editor: Andy Kleinschmidt Web Editor: Tom Rosati
Information presented above and where trade names are used, they are supplied with the understanding that no discrimination is intended and no endorsement by Ohio State University Extension is implied. Although every attempt is made to produce information that is complete, timely, and accurate, the pesticide user bears responsibility of consulting the pesticide label and adhering to those directions.
All educational programs conducted by Ohio State University Extension are available to clientele on a nondiscriminatory basis without regard to race, color, creed, religion, sexual orientation, national origin, gender, age, disability or Vietnam-era veteran status.
Issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture, Keith L. Smith, Director, Ohio State University Extension.
TDD # 1 (800) 589-8292 (Ohio only) or (614) 292-1868
| C.O.R.N. | Newsletter | Archive | Search | Questions? | Ohioline | Publications |