The research was supported by the National Research Council (CNR) of Italy. The authors thank GIRE members for collecting seed samples and are grateful to Alison Garside for revising the English.

1. Seed Sampling and Storage
<ol>
	<li>Monitor cultivated fields for unjustified poor herbicide performance, i.e., not due to adverse climatic conditions or low quality herbicide treatments.</li>
	<li>Collect a seed sample from one species at a time and assign a unique code. Mature seeds are usually collected before crop harvesting from plants that had survived the herbicide treatment(s). Timely monitor to observe if the seeds are shed by mother plant when mature.</li>
	<li>Fill in a form for each sample indicating the assigned unique code, name of species, collection date, GPS coordinates, municipality, farmer&#8217;s name, field size, infestation level, crop, herbicide(s) used during the season and historical records of the field.</li>
	<li>Collect seeds from at least 30 randomly selected plants that are representative of the field infestation. Ensure that the seed sample contains at least 5,000 mature seeds. For an obligate outcrossing weed species (e.g., Lolium spp. or Amaranthus spp.), reduce the number of plants to 10-15, keeping the total number of seeds around 5,000 11.</li>
	<li>Sub-sample the field if patches of weeds are scattered over large areas (more than a hectare) as different herbicide resistant biotypes are selected.</li>
	<li>Store seeds in unsealed paper bags labeled with the unique code assigned.</li>
	<li>Allow moisture to evaporate but do not expose seeds to high temperature (i.e., avoid leaving them in a car under the sun) or to extreme temperature fluctuations to avoid induction of secondary dormancy.</li>
	<li>Clean (remove chaff, de-hull seeds, etc.) and store them at ambient temperature in a dry room. After conducting the first resistance tests, store the seeds for long periods of time in a dark room at 4 &#176;C, preferably in vacuum-sealed plastic bags. In this way seeds preserve their viability for a significantly longer time.</li>
</ol>
2. Seed Dormancy Breaking
NOTE: Seed dormancy provides a flexible and efficient mechanism that enables weeds to adapt and persist in agro-ecosystems. To break dormancy and allow seed germination, different protocols have to be used depending on weed species, i.e., the type of dormancy 16. 
There are three main ways to remove dormancy:
<ol>
	<li>Vernalization 
	NOTE: To obtain simultaneous germination and seedling emergence, a period of seed vernalization ranging from a few days to a week is required to remove physiological dormancy from many species: e.g., Amaranthus retroflexus, Chenopodium album, Lolium spp., Avena fatua, Polygonum persicaria, Phalaris paradoxa17-19. A longer period of up to 15 days is required for Papaver rhoeas, Cyperus difformis and Ammania coccinea and up to 30 days for Schoenoplectus mucronatus20.
	<ol>
		<li>Put some deionized water in plastic dishes. Cut two layers of filter paper and soak them in the water, remove any excess. Place the air-dried seeds on the paper. Transfer the plastic dishes to a fridge at 4 &#176;C for the required period of time.</li>
	</ol>
	</li>
	<li>Scarification 
	NOTE: Some weed species are more recalcitrant to germination than others due to mechanical dormancy, i.e. seed coat characteristics, and necessitate the use of a chemical scarification using sulfuric acid to germinate 21.
	<ol>
		<li>Prepare a beaker with concentrated sulfuric acid (95-98%). Prepare a beaker full of water. Put the seeds in an envelope of non-woven fabric.</li>
		<li>Soak e.g., Echinochloa spp. or Sorghum halepense seeds for 20 min or 5 min, respectively, in concentrated sulfuric acid.</li>
		<li>Take the envelope out of the beaker using a pair of tweezers and put it in the beaker full of water. Open the envelope, put the seeds in a small colander and rinse them thoroughly under running water.</li>
	</ol>
	Variant: A. plantago-aquatica needs a different protocol 22, 23
	<ol>
		<li>Soak the seeds for 2 min in chloroform. Rinse the seeds with deionized water and dry them with absorbent paper. Dunk the seeds in 80% sulfuric acid for 5 min.</li>
		<li>Put the seeds in a small colander and rinse them thoroughly under running water.</li>
	</ol>
	</li>
	<li>Post-harvest seed maturation 
	NOTE: Seeds of other weed species do not germinate at all for a few months after maturity, regardless of the method used to break dormancy.
	<ol>
		<li>Store the seeds for a period of at least 3-4 months at RT and low humidity and then follow the above protocols for dormancy breaking (e.g., Oryza sativa var. sylvatica or P. rhoeas).</li>
	</ol>
	</li>
</ol>
3. Seed Germination
<ol>
	<li>Place seeds to be germinated in plastic dishes containing 0.6% (m/v) agar with 0.1% potassium nitrate (KNO3) added:
	<ol>
		<li>Prepare a solution of agar at 0.6% + 0.1% KNO3 using deionized water. Dissolve the agar in a microwave oven.</li>
		<li>Pour the agar solution into plastic dishes. Cool the substrate and then put in the seeds.</li>
	</ol>
	Variant: Soil can also be used as a substrate in plastic dishes. This practice is particularly efficient for Echinochloa spp., S. halepense and Lolium spp.</li>
	<li>Place plastic dishes in a germination cabinet for about a week with light and temperature conditions depending on the optimum conditions for each weed species. For most winter species, the temperature range is 15/25 &#176;C night/day and 12 hr photoperiod with neon tubes providing a Photosynthetic Photon Flux Density (PPFD) of 15-30 &#181;mol m-2 sec-1. For many summer species, the temperature range is 15/30 &#176;C night/day. 
	Variant: Some species, such as S. halepense, need a heat treatment. Therefore after the scarification, seeds of S. halepense are subjected to the following conditions: cycles of 4 hr at 45 &#176;C and 20 hr at 24 &#176;C for three days in the germination cabinet, and then three days in normal conditions.</li>
</ol>
4. Seedling Transplanting and Growth
<ol>
	<li>Transplant fifteen to twenty seedlings into plastic trays (325 x 265 x 95 mm) filled with a standard potting mix (60% silty loam soil, 15% sand, 15% agriperlite and 10% peat &#8212; by volume). 
	NOTE: Transplanting, instead of direct sowing, allows a uniform stand of plants at the same growth stage to be obtained, which is an important pre-condition to optimize the performance of the herbicide treatment.</li>
	<li>Identify each tray with a barcode including all information for the unique identification: population code, herbicide being tested, replicate number and progressive tray number.</li>
	<li>Place trays in a heated greenhouse and water plants as needed to maintain the substrate at or near field capacity. 
	NOTE: The growth temperature varies depending on the weed species. Often tests are done during autumn/winter/spring, so light is supplemented using 400 W metal-halide lamps, which provide a PPFD of about 150 &#181;mol m-2 sec-1 and a 12 hr photoperiod 24, 19. Summer weed species with C4 photosynthetic cycle usually require higher light intensity and therefore tests are done in late spring-summer or the supplemented light intensity is about 400 &#181;mol m-2 sec-1 with a 14 hr photoperiod.</li>
	<li>Use a different protocol for some weed species infesting paddy rice, e.g., A. plantago-aquatica, S. mucronatus and C. difformis as described in 22.
	<ol>
		<li>Transplant the seedlings into polystyrene trays with 24 round cells (55 mm diameter, 64 mm deep) filled with 60% silty loam soil, 30% sand and 10% peat (by volume).</li>
		<li>Set the trays in 12 cm deep plastic containers filled with water and battened down by screwed stainless steel rods to prevent them floating (Figure 1).</li>
		<li>Maintain water level in the containers at 1-2 cm below the level of the soil surface and add 1.5 g of copper sulfate to each container (which contains 10-12 L of water) to avoid the proliferation of algae.</li>
	</ol>
	</li>
</ol>
5. Herbicide Treatments
<ol>
	<li>Treatments with pre-emergence herbicides:
	<ol>
		<li>After about three days in the germination cabinet as described in section 3, transplant the germinating seeds into plastic trays containing the substrate described above and cover with a layer of soil (about 1 cm). This is a critical step in order to ensure that seedlings will not emerge due to the herbicide effect rather than an excessive burial depth.</li>
		<li>Take the substrate to field capacity by placing the trays, which have a few holes at the bottom, on saucers filled with water.</li>
		<li>One day after transplanting, treat the trays with the pre-emergence herbicide 25.</li>
		<li>Keep the substrate at or near field capacity by adding water as needed both from above and below by capillarity from the saucer. This procedure favors the permanence of the herbicide at the proper depth (i.e., where the germinating seeds are) for a good treatment efficacy.</li>
	</ol>
	</li>
	<li>Treatments with post-emergence herbicides:
	<ol>
		<li>Spray plants when they reach the 2-3 leaf stage (i.e., growth stage 12-13 of the Extended BBCH growth scale 26).</li>
		<li>Starting from the day after the treatment, set the irrigation system according to the water requirements of the weed species and the season (e.g., for Echinochloa spp. it delivers water for 3 min 4 times per day, at regular intervals from 9 am to 9 pm). Water is distributed using an automatic sprinkler irrigation system. 
		Variant: glyphosate is applied at plant stage BBCH 14&#8211;21.</li>
	</ol>
	</li>
	<li>Herbicide preparation and distribution. 
	NOTE: All herbicides (pre- and post-emergence) are applied as commercial formulations with recommended surfactants at two doses, recommended field dose (1x) and three times that (3x).
	<ol>
		<li>If needed, prepare the surfactant solution in bulk according to the label instructions; the final concentration is usually expressed as a percentage of the final volume (e.g., 0.3%) or as volume to be distributed per unit area (e.g., 1 L ha-1).</li>
		<li>Use the surfactant solution as solvent for the herbicide (solute) solution in order to keep the right concentration of the active ingredient. Prepare the most concentrated herbicide solution first (3x). Calculate the quantity of commercial product to be dissolved in the surfactant solution (or in deionized water if a surfactant is not necessary) using the following equation: 
		Doseherb = [(Dosefield x Dosemax) x Vfin] / Vdel 
		Where: Doseherb = herbicide dose (ml), Dosefield = herbicide field dose (ml ha-1), Dosemax = maximum dose delivered, Vfin = final volume of the solution (L), Vdel = volume delivered by the bench sprayer (L ha-1).</li>
		<li>Dilute (2:1, v/v) the herbicide solution 3x to prepare the less concentrated one (1x). This procedure reduces the chance of making mistakes when weighing or pipetting the herbicides. Herbicide solution concentration is expressed as volume to be distributed per unit area (L ha-1).</li>
		<li>Start the sequence of treatment with the lower herbicide dose (1x). In this way there is no need to wash the spraying cabinet between two treatments with the same herbicide.</li>
		<li>Distribute the herbicide solution using a precision bench sprayer delivering 300 L ha-1 (&#177;1%), at a pressure of 215 kPa, and a speed of 0.75 m sec-1, with a boom equipped with three flat-fan (extended range) hydraulic nozzles.</li>
		<li>Wash the spraying cabinet twice when the herbicide is changed using bleach 1% (v/v) and then rinse. 
		Variant: glyphosate is applied with a spray volume of 200 L ha-1 27. 
		NOTE: Particular attention has to be paid when highly biological herbicides, such the sulfonylureas sulfometuron or flazasulfuron, are used. In the latter case do one washing with a bleach solution and another with ammonia (2.5% v/v), followed by a careful rinse with water.</li>
	</ol>
	</li>
</ol>
6. Collection and Analysis of the Data
<ol>
	<li>Through a barcode reader, which automatically identifies each tray, record the number of plants that survived the treatment as well as the Visual Estimated Biomass (VEB). Plants are assessed as being dead if they show no active growth regardless of color or other appearance.
	<ol>
		<li>Make the assessment three or four weeks after treatment (WAT) depending on the herbicides tested (e.g., three WAT for ACCase inhibitors and four WAT for ALS inhibitors or glyphosate).</li>
		<li>Evaluate the general treatment efficacy by including a susceptible population (check S) in all experiments, i.e., a population collected in a site which was never or seldom treated with herbicides.</li>
		<li>Express plant survival as percentage of the number of treated plants, counted just before the herbicide treatment, and calculate the standard error (S.E.) per mean value (mean value of the two replicates).</li>
		<li>The VEB is obtained through a visual comparison of plant biomass between treated and not-treated check of the same population 25, 28. A score, ranging from 10 for plants not affected by the herbicide (compared with the not-treated check) to 0 when the plants are clearly dead, is given to each treated tray.</li>
	</ol>
	</li>
	<li>Ascribe populations to four categories based on the results obtained from treatments with two herbicide doses: S when less than 5% of plants survived the herbicide dose 1x, SR when survivors ranged from 5% to 20% at herbicide dose 1x, R when more than 20% of plants survived the herbicide dose 1x and RR when survivors are more than 20% at herbicide dose 1x and more than 10% at herbicide dose 3x 17.</li>
</ol>

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The authors declare that they have no competing financial interests.

Several steps within the protocols are critical for a successful assessment of herbicide resistance in a population: 1) seeds should be collected when mature from plants that had survived the herbicide treatment(s). Maturation of the seeds on the mother plant is crucial to avoid difficulties in seed germination later; 2) the proper storage of seeds is recommended to avoid proliferation of molds that would prevent germination; 3) seedlings should be treated at the right growth stage, as reported on the label of the herbic...

Herbicides are the most extensively used weed control measure, accounting for up to 50% of the global plant protection market 1. They are relatively cheap tools, avoid labor-intensive and time-consuming soil cultivation practices, and ultimately result in cost-effective, safe and profitable food production 2. However, the great phenological and genetic variability present in many weed species, together with an over-reliance on herbicide use, frequently results in the selection of herbicide-resistant weed populations. The introduction of selective herbicides with a very specific metabolic target 3-5 has dramatically increased the number of resistance cases over the years. To date, 240 weed species (140 dicots and 100 monocots) worldwide have evolved resistance to different herbicide Sites of Action (SoA) 4. This is a major concern for weed management and more in general for sustainable crop production.
Early detection of resistance, based on reliable tests, frequently performed in a greenhouse, is a key step to manage herbicide resistant weeds. Different approaches have been developed according to the aims, required level of accuracy, time and resources available, as well as the weed species considered 6-12. However, when confirmation of the resistance status of a new weed biotype is required (i.e., a group of individuals that share several physiological characteristics, including the ability to survive one or more herbicides belonging to a particular group used at a dose that would normally control them), a robust whole plant bioassay needs to be performed in a controlled environment 4, 11.
A biotype is seldom resistant to just one herbicide. Each biotype is therefore characterized by a certain resistance pattern, i.e., number and type of SoA of the herbicides it is resistant to, and by a given resistance level to each herbicide 13. The early and reliable determination of the pattern of cross or multiple resistance 5, 14 is important for field resistance management.
It is worth mentioning that herbicide resistance has nothing to do with the natural tolerance that some weed species exhibit towards some herbicides, e.g., dicot species vs. ACCase-inhibiting herbicides, monocot species vs. 2,4-D, Equisetum arvense vs. glyphosate.
This paper presents a robust approach for testing putative herbicide resistant biotypes sampled in fields where poor control by herbicide(s) had been reported. Relevant variants to the standard protocols in relation to the weed species involved are presented. The advantages over alternative techniques/protocols based on either whole plant bioassays using only one herbicide dose 15,or treating seeds in Petri dishes 8 are related to the higher reliability and the possibility of inferring the resistance level because of the inclusion of two herbicide doses in the experiments. However, for routine resistance testing, the same methods can be applied at only one herbicide dose, so reducing the costs.
As well as allowing confirmation of the resistance status, the information obtained can be used for both optimizing the following research steps and/or devising sound resistance management strategies.

<table><tbody><tr><td>Paper bags</td><td>Celcar SAS</td><td></td><td></td></tr><tr><td>Plastic dishes</td><td>ISI plast S.p.A.</td><td>SO600</td><td>Transparent plastic</td></tr><tr><td>Sulfuric acid 95-98%</td><td>Sigma-Aldrich</td><td>320501</td><td></td></tr><tr><td>Non-woven fabric</td><td>Carretta Tessitura</td><td>Art.TNT17</td><td>Weight 17 g m&lt;sup&gt;-2&lt;/sup&gt;</td></tr><tr><td>Chloroform &gt;99.5%</td><td>Sigma-Aldrich</td><td>C2432</td><td></td></tr><tr><td>Agar</td><td>Sigma-Aldrich</td><td>A1296</td><td></td></tr><tr><td>Potassium nitrate &gt;99.0%</td><td>Sigma-Aldrich</td><td>P8394</td><td></td></tr><tr><td>Plastic containers</td><td>Giganplast</td><td>1875/M</td><td>600 x 400 x 110 mm</td></tr><tr><td>Plastic trays</td><td>Piber plast</td><td>G1210A</td><td>325 x 265 x 95 mm</td></tr><tr><td>Polystyrene trays</td><td>Plastisavio</td><td>S24</td><td>537 x 328 x 72 mm, 24 round cells (6x4)</td></tr><tr><td>Copper sulfate</td><td>Sigma-Aldrich</td><td>451657</td><td></td></tr><tr><td>Agriperlite</td><td>Blu Agroingross sas</td><td>AGRI100</td><td></td></tr><tr><td>Peat</td><td>Blu Agroingross sas</td><td>TORBA250</td><td></td></tr><tr><td>Germination cabinet</td><td>KW</td><td>W87R</td><td></td></tr><tr><td>Nozzles</td><td>Teejet</td><td>XR11002-VK, TP11001-VH</td><td>The second type of nozzles are used only for glyphosate</td></tr><tr><td>Barcode generator</td><td>Toshiba TEC</td><td>SX4</td><td></td></tr><tr><td>Labels with barcode</td><td>Felga</td><td>TT20200</td><td>Stick-in labels with rounded corners</td></tr><tr><td>Barcode reader</td><td>Cipherlab</td><td>8300-L</td><td>Portable data terminal</td></tr><tr><td>Bench sprayer</td><td></td><td></td><td>Built in house</td></tr><tr><td colspan="4">&lt;strong&gt;Herbicides included in the results:&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Commercial product&lt;/strong&gt;</td><td>&lt;strong&gt;Active ingredient&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>Altorex</td><td>imazamox</td><td>BASF</td><td></td></tr><tr><td>Azimut</td><td>florasulam</td><td>Dow AgroSciences</td><td></td></tr><tr><td>Biopower</td><td></td><td>Bayer Crop Science</td><td>Surfact to be used with Hussar WG</td></tr><tr><td>Dash</td><td></td><td>BASF</td><td>Surfact to be used with Altorex</td></tr><tr><td>Granstar</td><td>tribenuron-methyl</td><td>Dupont</td><td></td></tr><tr><td>Gulliver</td><td>azimsulfuron</td><td>Dupont</td><td></td></tr><tr><td>Hussar WG</td><td>iodosulfuron</td><td>Bayer Crop Science</td><td></td></tr><tr><td>Nominee</td><td>bispyribac-Na</td><td>Bayer Crop Science</td><td></td></tr><tr><td>Roundup</td><td>glyphosate</td><td>Monsanto</td><td></td></tr><tr><td>Trend</td><td></td><td>Dupont</td><td>Surfact to be used with Granstar and Gulliver</td></tr><tr><td>Viper</td><td>penoxsulam</td><td>Dow AgroSciences</td><td></td></tr><tr><td>Weedone LV4</td><td>2,4-D</td><td>Isagro</td><td></td></tr></tbody></table>

protocols for robust herbicide resistance testing in different weed species

Robust protocols to test putative herbicide resistant weed populations at whole plant level are essential to confirm the resistance status. The presented protocols, based on whole-plant bioassays performed in a greenhouse, can be readily adapted to a wide range of weed species and herbicides through appropriate variants. Seed samples from plants that survived a field herbicide treatment are collected and stored dry at low temperature until used. Germination methods differ according to weed species and seed dormancy type. Seedlings at similar growth stage are transplanted and maintained in the greenhouse under appropriate conditions until plants have reached the right growth stage for herbicide treatment. Accuracy is required to prepare the herbicide solution to avoid unverifiable mistakes. Other critical steps such as the application volume and spray speed are also evaluated. The advantages of this protocol, compared to others based on whole plant bioassays using one herbicide dose, are related to the higher reliability and the possibility of inferring the resistance level. Quicker and less expensive in vivo or in vitro diagnostic screening tests have been proposed (Petri dish bioassays, spectrophotometric tests), but they provide only qualitative information and their widespread use is hindered by the laborious set-up that some species may require. For routine resistance testing, the proposed whole plant bioassay can be applied at only one herbicide dose, so reducing the costs.

To assess the resistance status of a putative resistant population, it is fundamental to include a susceptible check in the assay in order to verify the herbicide efficacy. The results of a screening test conducted on P. rhoeas populations, a weed infesting wheat fields, are reported in Figure 2, where the efficacy of four post-emergence herbicides on a susceptible check (09-36) and on the suspected resistant one (10-91) are presented. Population 09-36 was completely controlled by the ALS inhibi...

Watch this Scientific Journal Video about Protocols for Robust Herbicide Resistance Testing in Different Weed Species at JoVE.com

Protocols for Robust Herbicide Resistance Testing in Different Weed Species

A robust and flexible approach to confirm herbicide resistance in weed populations is presented. This protocol allows the herbicide resistance levels to be inferred and applied to a wide range of weed species and herbicides with minor adaptations.

Robust protocols to test putative herbicide resistant weed populations at whole plant level are essential to confirm the resistance status. The presented ...

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Research

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14.6K Views.  National Research Council (CNR), Italy. A robust and flexible approach to confirm herbicide resistance in weed populations is presented. This protocol allows the herbicide resistance levels to be inferred and applied to a wide range of weed species and herbicides with minor adaptations.

Video: Protocols for Robust Herbicide Resistance Testing in Different Weed Species

Measuring Rates of Herbicide Metabolism in Dicot Weeds with an Excised Leaf Assay

In order to isolate and accurately determine rates of herbicide metabolism in an obligate-outcrossing dicot weed, waterhemp (Amaranthus tuberculatus), we developed an excised leaf assay combined with a vegetative cloning strategy to normalize herbicide uptake and remove translocation as contributing factors in herbicide-resistant (R) and &#8211;sensitive (S) waterhemp populations. Biokinetic analyses of organic pesticides in plants typically include the determination of uptake, translocation (delivery to the target site), metabolic fate, and interactions with the target site. Herbicide metabolism is an important parameter to measure in herbicide-resistant weeds and herbicide-tolerant crops, and is typically accomplished with whole-plant tests using radiolabeled herbicides. However, one difficulty with interpreting biokinetic parameters derived from whole-plant methods is that translocation is often affected by rates of herbicide metabolism, since polar metabolites are usually not mobile within the plant following herbicide detoxification reactions. Advantages of the protocol described in this manuscript include reproducible, accurate, and rapid determination of herbicide degradation rates in R and S populations, a substantial decrease in the amount of radiolabeled herbicide consumed, a large reduction in radiolabeled plant materials requiring further handling and disposal, and the ability to perform radiolabeled herbicide experiments in the lab or growth chamber instead of a greenhouse. As herbicide resistance continues to develop and spread in dicot weed populations worldwide, the excised leaf assay method developed and described herein will provide an invaluable technique for investigating non-target site-based resistance due to enhanced rates of herbicide metabolism and detoxification.

This manuscript describes how herbicide metabolism rates can be effectively quantified with excised leaves from a dicot weed, thereby reducing variability and removing any possible confounding effects of herbicide uptake or translocation typically observed in whole-plant assays.

In order to isolate and accurately determine rates of herbicide metabolism in an obligate-outcrossing dicot weed, waterhemp (Amaranthus tuberculatus), we ...

Protocols for Quantifying Transferable Pesticide Residues in Turfgrass Systems

Plant canopies in established turfgrass systems can intercept an appreciable amount of sprayed pesticides, which can be transferred through various routes onto humans. For this reason, transferable pesticide residue experiments are required for registration and re-registration by the United States Environmental Protection Agency (USEPA). Although such experiments are required, limited specificity is required pertaining to experimental approach. Experimental approaches used to assess pesticide transfer to humans including hand wiping with cotton gloves, modified California roller (moving a roller of known mass over cotton cloth) and soccer ball roll (ball wrapped with sorbent strip) over three treated turfgrass species (creeping bentgrass, hybrid bermudagrass and tall fescue maintained at 0.4, 5 and 9 cm, respectively) are presented. The modified California roller is the most extensively utilized approach to date, and is best suited for use at low mowing heights due to its reproducibility and large sampling area. The soccer ball roll is a less aggressive transfer approach; however, it mimics a very common occurrence in the most popular international sport, and has many implications for nondietary pesticide exposure from hand-to-mouth contact. Additionally, this approach may be adjusted for other athletic activities with limited modification. Hand wiping is the best approach to transfer pesticides at higher mowing heights, as roller-based approaches can lay blades over; however, it is more subjective due to more variable sampling pressure. Utility of these methods across turfgrass species is presented, and additional considerations to conduct transferable pesticide residue research in turfgrass systems are discussed.

Transferable pesticide residue experiments in turfgrass systems are integral components of human risk exposure assessments. Experimental approaches to measure transferable residues should be adjusted to the human interaction of interest and turfgrass system dynamics. Three transferable pesticide residue protocols are presented and the suitability across three turfgrass systems is discussed.

Plant canopies in established turfgrass systems can intercept an appreciable amount of sprayed pesticides, which can be transferred through various routes ...

Screening Cotton Genotypes for Reniform Nematode Resistance

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium hirsutum) varieties to improve nematode management. Most protocols involve extracting vermiform nematodes or eggs from the cotton root system or potting soil to determine population density or reproduction rate. These approaches are generally time-consuming with a small number of genotypes evaluated. An alternative approach is described here in which the root system is visually examined for nematode infection. The protocol involves inoculating cotton seedling 7 days after planting with vermiform nematodes and determining the number of females attached to the root system 28 days after inoculation. Data are expressed as the number of females per gram of fresh root weight to adjust for variation in root growth. The protocol provides an excellent method for evaluating host-plant resistance associated with the ability of the nematode to establish an infection site; however, resistance that hinders nematode reproduction is not assessed. As with other screening protocols, variation is commonly observed in nematode infection among individual genotypes within and between experiments. Data are presented to illustrate the range of variation observed using the protocol. To adjust for this variation, control genotypes are included in experiments. Nonetheless, the protocol provides a simple and rapid method to evaluate host-plant resistance. The protocol has been successfully used to identify resistant accessions from the G. arboreum germplasm collection and evaluate segregating populations of more than 300 individuals to determine the genetics of resistance. A vegetative propagation method for recovering plants for resistance breeding was also developed. After removal of the root system for nematode evaluation, the vegetative shoot is replanted to allow the development of a new root system. More than 95% of the shoots typically develop a new root system with plants reaching maturity.

Here, a protocol is presented for the rapid non-destructive screening of cotton genotypes for reniform nematode resistance. The protocol involves visually examining the roots of nematode-infected cotton seedlings to determine infection response. The vegetative shoot from each plant is then propagated to recover plants for seed production.

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium ...

Genetics

Quantification of the Potential Impact of Glyphosate-Based Products on Microbiomes

Glyphosate-based products (GBP) are the most common broad-spectrum herbicides worldwide. The target of glyphosate is the enzyme 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) in the shikimate pathway, which is virtually universal in plants. The inhibition of the enzyme stops the production of three essential amino acids: phenylalanine, tyrosine, and tryptophan. EPSPS is also present in fungi and prokaryotes, such as archaea and bacteria; thus, the use of GBP may have an impact on the microbiome composition of soils, plants, herbivores, and secondary consumers. This article aims to present general guidelines to assess the effect of GBP on microbiomes from field experiments to bioinformatics analyses and provide a few testable hypotheses. Two field experiments are presented to test the GBP on non-target organisms. First, plant-associated microbes from 10 replicated control and GBP treatment plots simulating no-till cropping are sampled and analyzed. In the second experiment, samples from experimental plots fertilized by either poultry manure containing glyphosate residues or non-treated control manure were obtained. Bioinformatics analysis of EPSPS protein sequences is utilized to determine the potential sensitivity of microbes to glyphosate. The first step in estimating the effect of GBP on microbiomes is to determine their potential sensitivity to the target enzyme (EPSPS). Microbial sequences can be obtained either from public repositories or by means of PCR amplification. However, in the majority of field studies, microbiome composition has been determined based on universal DNA markers such as the 16S rRNA and the internal transcribed spacer (ITS). In these cases, sensitivity to glyphosate can only be estimated through a probabilistic analysis of EPSPS sequences using closely related species. The quantification of the potential sensitivity of organisms to glyphosate, based on the EPSPS enzyme, provides a robust approach for further experiments to study target and non-target resistant mechanisms.

Glyphosate-based products (GBP) are the most common broad-spectrum herbicides worldwide. In this article, we introduce general guidelines to quantify the effect of GBP on microbiomes, from field experiments to bioinformatics analyses.

Glyphosate-based products (GBP) are the most common broad-spectrum herbicides worldwide. The target of glyphosate is the enzyme ...

Bioassays for Monitoring Insecticide Resistance

Pest resistance to pesticides is an increasing problem because pesticides are an integral part of high-yielding production agriculture. When few products are labeled for an individual pest within a particular crop system, chemical control options are limited. Therefore, the same product(s) are used repeatedly and continual selection pressure is placed on the target pest. There are both financial and environmental costs associated with the development of resistant populations. The cost of pesticide resistance has been estimated at approximately $ 1.5 billion annually in the United States. This paper will describe protocols, currently used to monitor arthropod (specifically insects) populations for the development of resistance. The adult vial test is used to measure the toxicity to contact insecticides and a modification of this test is used for plant-systemic insecticides. In these bioassays, insects are exposed to technical grade insecticide and responses (mortality) recorded at a specific post-exposure interval. The mortality data are subjected to Log Dose probit analysis to generate estimates of a lethal concentration that provides mortality to 50% (LC50) of the target populations and a series of confidence limits (CL's) as estimates of data variability. When these data are collected for a range of insecticide-susceptible populations, the LC50 can be used as baseline data for future monitoring purposes. After populations have been exposed to products, the results can be compared to a previously determined LC50 using the same methodology.

This manuscript demonstrates and discusses techniques used to survey pesticide susceptibility and detect resistance to contact and systemic pesticides in arthropod pests.

Pest resistance to pesticides is an increasing problem because pesticides are an integral part of high-yielding production agriculture. When few products ...

Biology

Implementation of In Vitro Drug Resistance Assays: Maximizing the Potential for Uncovering Clinically Relevant Resistance Mechanisms

Although targeted therapies are initially effective, resistance inevitably emerges. Several methods, such as genetic analysis of resistant clinical specimens, have been applied to uncover these resistance mechanisms to facilitate follow-up care. Although these approaches have led to clinically relevant discoveries, difficulties in attaining the relevant patient material or in deconvoluting the genomic data collected from these specimens have severely hampered the path towards a cure. To this end, we here describe a tool for expeditious discovery that may guide improvement in first-line therapies and alternative clinical management strategies. By coupling preclinical in vitro or in vivo drug selection with next-generation sequencing, it is possible to identify genomic structural variations and/or gene expression alterations that may serve as functional drivers of resistance. This approach facilitates the spontaneous emergence of alterations, enhancing the probability that these mechanisms may be observed in the patients. In this protocol we provide guidelines to maximize the potential for uncovering single nucleotide variants that drive resistance using adherent lines.

Implementation of In Vitro Drug Resistance Assays: Maximizing the Potential for Uncovering Clinically Relevant Resistance Mechanisms

Drug resistance to targeted therapeutics is widespread and the need to identify mechanisms of resistance--prior to or following clinical onset--is critical for guiding alternative clinical management strategies. Here, we present a protocol to couple derivation of drug-resistant lines in vitro with sequencing to expedite discovery of these mechanisms.

Although targeted therapies are initially effective, resistance inevitably emerges. Several methods, such as genetic analysis of resistant clinical ...

Medicine

Characterizing Herbivore Resistance Mechanisms: Spittlebugs on Brachiaria spp. as an Example

Plants can resist herbivore damage through three broad mechanisms: antixenosis, antibiosis and tolerance1. Antixenosis is the degree to which the plant is avoided when the herbivore is able to select other plants2. Antibiosis is the degree to which the plant affects the fitness of the herbivore feeding on it1.Tolerance is the degree to which the plant can withstand or repair damage caused by the herbivore, without compromising the herbivore's growth and reproduction1. The durability of herbivore resistance in an agricultural setting depends to a great extent on the resistance mechanism favored during crop breeding efforts3.
We demonstrate a no-choice experiment designed to estimate the relative contributions of antibiosis and tolerance to spittlebug resistance in Brachiaria spp. Several species of African grasses of the genus Brachiaria are valuable forage and pasture plants in the Neotropics, but they can be severely challenged by several native species of spittlebugs (Hemiptera: Cercopidae)4.To assess their resistance to spittlebugs, plants are vegetatively-propagated by stem cuttings and allowed to grow for approximately one month, allowing the growth of superficial roots on which spittlebugs can feed. At that point, each test plant is individually challenged with six spittlebug eggs near hatching. Infestations are allowed to progress for one month before evaluating plant damage and insect survival. Scoring plant damage provides an estimate of tolerance while scoring insect survival provides an estimate of antibiosis. This protocol has facilitated our plant breeding objective to enhance spittlebug resistance in commercial brachiariagrases5.

Characterizing Herbivore Resistance Mechanisms: Spittlebugs on Brachiaria spp. as an Example

This video explains mechanisms of host plant resistance to herbivory and demonstrates a no-choice test that estimates the relative contributions of antibiosis and tolerance to spittlebug resistance in Brachiaria spp.

Plants can resist herbivore damage through three broad mechanisms: antixenosis, antibiosis and tolerance1. Antixenosis is the degree to which the plant is ...

A PCR-based Genotyping Method to Distinguish Between Wild-type and Ornamental Varieties of Imperata cylindrica

Wild-type I. cylindrica (cogongrass) is one of the top ten worst invasive plants in the world, negatively impacting agricultural and natural resources in 73 different countries throughout Africa, Asia, Europe, New Zealand, Oceania and the Americas1-2. Cogongrass forms rapidly-spreading, monodominant stands that displace a large variety of native plant species and in turn threaten the native animals that depend on the displaced native plant species for forage and shelter. To add to the problem, an ornamental variety [I. cylindrica var. koenigii (Retzius)] is widely marketed under the names of Imperata cylindrica 'Rubra', Red Baron, and Japanese blood grass (JBG). This variety is putatively sterile and noninvasive and is considered a desirable ornamental for its red-colored leaves. However, under the correct conditions, JBG can produce viable seed (Carol Holko, 2009 personal communication) and can revert to a green invasive form that is often indistinguishable from cogongrass as it takes on the distinguishing characteristics of the wild-type invasive variety4 (Figure 1). This makes identification using morphology a difficult task even for well-trained plant taxonomists. Reversion of JBG to an aggressive green phenotype is also not a rare occurrence. Using sequence comparisons of coding and variable regions in both nuclear and chloroplast DNA, we have confirmed that JBG has reverted to the green invasive within the states of Maryland, South Carolina, and Missouri. JBG has been sold and planted in just about every state in the continental U.S. where there is not an active cogongrass infestation. The extent of the revert problem in not well understood because reverted plants are undocumented and often destroyed.
Application of this molecular protocol provides a method to identify JBG reverts and can help keep these varieties from co-occurring and possibly hybridizing. Cogongrass is an obligate outcrosser and, when crossed with a different genotype, can produce viable wind-dispersed seeds that spread cogongrass over wide distances5-7. JBG has a slightly different genotype than cogongrass and may be able to form viable hybrids with cogongrass. To add to the problem, JBG is more cold and shade tolerant than cogongrass8-10, and gene flow between these two varieties is likely to generate hybrids that are more aggressive, shade tolerant, and cold hardy than wild-type cogongrass. While wild-type cogongrass currently infests over 490 million hectares worldwide, in the Southeast U.S. it infests over 500,000 hectares and is capable of occupying most of the U.S. as it rapidly spreads northward due to its broad niche and geographic potential3,7,11. The potential of a genetic crossing is a serious concern for the USDA-APHIS Federal Noxious Week Program. Currently, the USDA-APHIS prohibits JBG in states where there are major cogongrass infestations (e.g., Florida, Alabama, Mississippi). However, preventing the two varieties from combining can prove more difficult as cogongrass and JBG expand their distributions. Furthermore, the distribution of the JBG revert is currently unknown and without the ability to identify these varieties through morphology, some cogongrass infestations may be the result of JBG reverts. Unfortunately, current molecular methods of identification typically rely on AFLP (Amplified Fragment Length Polymorphisms) and DNA sequencing, both of which are time consuming and costly. Here, we present the first cost-effective and reliable PCR-based molecular genotyping method to accurately distinguish between cogongrass and JBG revert.

A PCR-based Genotyping Method to Distinguish Between Wild-type and Ornamental Varieties of Imperata cylindrica

We provide a cost-effective and rapid molecular genotyping protocol that employs variety-specific PCR primers that target DNA sequence differences within the chloroplast trnL-F spacer region to differentiate between varieties of Imperata cylindrica (cogongrass) that cannot be distinguished by morphology alone. These varieties include the federally listed noxious weed, cogongrass and closely-related, wide-spread ornamental variety, I. cylindrica var. koenigii (Japanese blood grass).

Wild-type I. cylindrica (cogongrass) is one of the top ten worst invasive plants in the world, negatively impacting agricultural and natural resources in ...

Repeatable Stair-step Assay to Access the Allelopathic Potential of Weedy Rice (Oryza sativa ssp.)

Weed competition contributes significantly to yield losses in cropping systems worldwide. The evolution of resistance in many weed species to continuously applied herbicides has presented the need for additional management methods. Allelopathy is a physiological process that some plant species possess that provide the plant with an advantage over its neighbors. Allelopathic crop varieties would be equipped with the ability to suppress the growth of surrounding competitors, thus reducing potential yield loss due to weed interference. This paper focuses on the construction and operation of a stair-step assay used for the screening of the allelopathic potential of a donor species (Oryza sativa) against a receiver weed species (Echinochloa crus-galli) in a greenhouse setting. The structure described in this paper serves as a stand for the plant samples and incorporates a timed watering system for the accumulation and distribution of allelochemicals. Allelochemicals produced by the plant roots are allowed to flow downward through a series of four pots separately into a collection tank and recycled back to the top plant through electric pumps. This method of screening provides an avenue for the allelochemicals from the donor plant to reach receiver plants without any resource competition, thus allowing quantitative measurement of the allelopathic potential of the selected donor plant. The allelopathic potential is measurable through the height reduction of the receiver plants. Preliminary screening data for the effectiveness of this method demonstrated height reduction in the receiver species, barnyardgrass (E. crus-galli), and thus the presence of allelopathic residues from the donor plant, weedy rice (Oryza sativa).

Repeatable Stair-step Assay to Access the Allelopathic Potential of Weedy Rice Oryza sativa ssp.

Allelopathy has shown promise as a useful supplemental weed control strategy in cropping systems. To determine the allelopathic potential of a desired plant specimen, a stair-step screening method is provided.

Weed competition contributes significantly to yield losses in cropping systems worldwide. The evolution of resistance in many weed species to continuously ...

High-throughput, Robust and Highly Time-flexible Method for Surface Sterilization of Arabidopsis Seeds

Arabidopsis is by far the plant model species most widely used for functional studies. The surface sterilization of Arabidopsis seeds is a fundamental step required towards this end. Thus, it is paramount to establish high-throughput Arabidopsis seed surface sterilization methods to handle tens to hundreds of samples (e.g., transgenic lines, ecotypes, or mutants) at once. A seed surface sterilization method based on the efficient elimination of liquid in tubes with a homemade suction device constructed from a common vacuum pump is presented in this study. By dramatically reducing labor-intensive hands-on time with this method handling several hundreds of samples in one day is possible with little effort. Series time-course analyses further indicated a highly flexible time range of surface sterilization by maintaining high germination rates. This method could be easily adapted for surface sterilization of other kinds of small seeds with simple customization of the suction device according to the seed size, and the speed desired to eliminate the liquid.

A high-throughput protocol for the surface sterilization of Arabidopsisthaliana (Arabidopsis) seeds is provided, optimizing the liquid handling steps with a simple suction device constructed with a vacuum pump. Hundreds of seed samples can be surface-sterilized in one day.

Arabidopsis is by far the plant model species most widely used for functional studies. The surface sterilization of Arabidopsis seeds is a fundamental ...

Protocols for Robust Herbicide Resistance Testing in Different Weed Species

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