S -metolachlor is used to control/suppress yellow nutsedge, annual grasses and several broadleaf weeds in sweetpotato. However, a decline in storage root quality is suspected when excessive rainfall occurs within 24-h after application. A greenhouse study was conducted to determine the effect of S-metolachlor application timing on sweetpotato growth and development. S-metolachlor treatments (0 and 1 kg·ha-1) were applied over-the-top at 0, 5 and ten days after transplanting (DA T ) and a simulated rainfall treatment delivered 25 mm of rain, 51 mm·h-1 intensity, immediately after herbicide application. Plants were harvested at 5, 10, 15, 20 and 80 DA T . During the first four harvests, roots were scanned and analyzed with WinRHIZO-Pro image analysis system to estimate root number, length, volume, and surface area along with aboveground growth parameters. At the final harvest, plant growth and biomass components, and quality of storage roots were recorded. Plants treated with S-metolachlor on day 0 and 5 DAT were significantly less than those of 10 DA T and untreated control for all measured parameters for the initial 20 days of plant growth. Even though vine length, leaf number, stem biomass, and total storage roots were not different among the treatments at 80 DAT, all other plant components and total biomass production and leaf area development for plants treated at 0 and 5 DAT were significantly (P < 0.05) less than from those of 10 DAT and the untreated control. Marketable storage root conversion efficiency declined by 18 % and 16% for plants treated at 0 and 5 DAT, respectively, relative to the untreated check. These results indicate that delaying S-metolachlor application to 10 DAT will be less damaging to sweetpotato growth and development, particularly marketable storage roots and yield.
Effective weed management, particularly within the first six weeks after transplanting, is essential to optimize sweetpotato yield [
S-metolachlor, one of few herbicides registered for use in sweetpotato, has a 24(c) Special Local Needs registration to control or suppress yellow nutsedge, annual grasses, and several small-seeded broadleaf weeds in sweetpotato production systems. S-metolachlor is physically and chemically equivalent to metolachlor (a 1:1 mixture of R- and S-isomers) but requires use rates 35% lower than metolachlor due to increased activity at the site of action in susceptible plants [
Even though S-metolachlor is an effective herbicide, growers are reluctant to use it because misshaped storage roots have been attributed to its use when applications are made soon after transplanting and followed by moderate to heavy rainfall [
Transplanted sweetpotato vine tip cuttings (slips) produce adventitious roots, some of which develop into storage roots through the proliferation of cambial cells that form starch-accumulating parenchyma [
A greenhouse study was conducted at the Rodney Foil Plant Science Center, Mississippi State University, Mississippi (lat. 33˚28'N, long. 88˚47'W) to determine the influence of S-metolachlor (Dual Magnum®, Syngenta Crop Protection Inc., Greensboro, NC, USA) application timing on “Beauregard” sweetpotato, which is the major cultivar grown in Mississippi, USA. White polyvinyl chloride pots (20 cm diameter × 30 cm deep) with detachable blue polyethylene (plug) bottoms containing a 2 mm drainage hole were filled with 600 g of coarse gravel then sandy loam soil (71% sand, 23% clay, 5% silt and 1% OM) obtained by mixing sand and topsoil in a 3:1 v/v ratio. On 21 June 2013, pots were irrigated to soil field capacity, and a single four node Beauregard slip was transplanted into each pot with two nodes below the soil surface and two nodes above the soil surface. Nodes above the soil surface each contained one recently fully expanded leaf.
Treatments were a factorial of three S-metolachlor application timings [0, 5 and 10 d after transplanting (DAT)] by two rates (0 and 1 kg ai ha−1) by five harvest timings (5, 10, 15, 20 and 80 DAT). S-metolachlor was applied with a tractor-mounted compressed-air spraying system fitted with Teejet 8002 XR flat fan nozzles (Teejet Spraying Systems Co., Wheaton, IL) and calibrated to deliver 140 L·ha−1 at 166 kPa. After the application, all pots from the same application timing received 25 mm of simulated rainfall at an intensity of 5.1 m·h−1. The rainfall simulator was modeled after one described by Meyer and Harmon [
In the greenhouse, pots were arranged in a split-split plot design with application timing as the main-plot factor, application rate as the sub-plot factor and harvest timing as the sub-sub-plot factor. Each treatment was replicated five times.
All plants received Hoagland’s nutrient solution in irrigation water at 8:00, 12:00 and 16:00 h each day, to ensure optimum nutrient [
Net photosynthetic rate, stomatal conductance, and intercellular CO2 concentration of the uppermost recently fully expanded main-stem leaves were measured between 10:00 and 12:00 h using an open gas exchange system (LI-6400, LiCOR Inc., Lincoln, NE, USA) at 20 DAT. While measuring photosynthesis, PAR, provided by a 6400-02 LED light source, was set to 1500-µmol·m−2·s−1, temperature inside the leaf cuvette was set to 30˚C (average growing temperatures in the greenhouse during the experimental period), RH was adjusted to near ambient level (50%), and leaf chamber CO2 concentration was set to 400-µmol·mol−1. Fluorescence was measured with the built-in leaf chamber fluorometer, which uses two red LEDs, center wavelength about 630 nm and a detector. The software in the instrument provides data on the fluorescence parameters and calculates parameters such as PSII reaction centers under light (Fv'/Fm') (LI-6400 Photosynthesis system, LI-COR, Inc.).
At 20 DAT, five 39-mm2 discs, one each from five recently fully expanded main-stem leaves, were cut from every plant using a cork borer. The discs were placed into a vial containing four mL dimethyl sulfoxide and held at room temperature overnight in the dark. Absorbance of the extract at 470, 648 and 664 nm was recorded using a Bio-Rad UV/VIS spectrophotometer (Bio-Rad Laboratories, Hercules, CA, USA) and chlorophyll a and b and carotenoid concentrations were computed following the formula of Chappelle et al. [
At each harvest timing, total vine length was measured and leaf number counted for each plant. Plant components (vines, leaves, roots) were separated, and leaf area measured (Li-COR 3100 Leaf Area Meter, LiCOR Inc.). All plant parts were bagged separately, oven-dried at 80˚C for 72 h and weighed. At harvests 5, 10, 15, and 20 DAT, roots were gently washed with water on a 3-mm mesh hardware cloth to remove soil. Roots were then placed into transparent acrylic trays (30 cm wide × 40 cm long × 2 depth) containing ~1 cm of water and scanned to acquire digital images using a flatbed scanner optimized for root analysis (Epson Expression 11000XL, Regent Instruments, Montreal, QC, Canada). Images were acquired at a resolution of 800 dpi then analyzed with root analysis system software (WinRHIZO Pro, Version 2012b, Regent Instruments, Montreal, QC, Canada) for root volume, length, and surface area. At the final harvest (80 DAT), storage roots were separated into marketable and non-marketable, counted, weighed, then oven-dried as described previously. Marketable storage roots were those longer than 7.6 cm, greater than 2.5 cm diameter, firm, smooth, and well-shaped without any disease [
At 20 DAT, washed storage roots ≥ 10 mm long were immediately removed and fixed in formalin-acetic acid-alcohol. The samples were dehydrated in a graded tertiary butyl alcohol series and embedded in paraplast. Blocks were sectioned at 8 microns with a rotary microtome (AmericanOptical Corp., Scientific Instrument Div., Buffalo, NY, USA), and sections stained with toluidine blue. Digital micrographs were taken with a Motic AE2000 microscope equipped with a Canon EOS Rebel T3i/600D 18.0-megapixel camera (MartinMicroscope Co., Easley, SC, USA).
All data were subjected to analysis of variance using the General Linear Model procedure of the Statistical Analysis System [
Leaf photosynthetic rate of the untreated check was 31.7 μmol CO2 m−2∙s−1 and decreased 21, 19% and 12% when S-metolachlor was applied at 0, 5 and 10 DAT, respectively (
Application timing | Photosynthesis | Chl a | Chl b | Chl a and b | Carotenoids |
---|---|---|---|---|---|
μmol CO2 m−2∙s−1 | ————µg·cm−2——— | ||||
Untreated check | 31.7A | 29.61A | 17.29A | 46.91A | 8.76 A |
0 DAT | 25.0B | 26.81C | 15.00B | 41.81C | 7.81B |
5 DAT | 25.6B | 27.84B | 15.62B | 43.45B | 8.32B |
10 DAT | 27.8AB | 29.49A | 17.13A | 46.62A | 8.71A |
Means within columns followed by different letters are significantly different based on Fisher’s least significant difference mean separation test (P < 0.05).
system in isolated pea chloroplast at metolachlor concentrations of up to 50 ppm. These results indicate that non-stomatal and non-photochemical processes are the causative factors limiting photosynthesis under S-metolachlor application. Similar to our results, Obando [
Leaf chlorophyll (Chl) a and b, total chlorophyll, and carotenoid concentrations decreased 9% and 6%, 12% and 9%, 11% and 7%, and 11% and 5% compared to the untreated check in plants treated 0 and 5 DAT, respectively (
At 20 DAT, there was an interaction between S-metolachlor application timing and harvest timing for vine length, leaf number, leaf area and total biomass (p < 0.05) (
Source of variation | Measured parameters | ||||||||
---|---|---|---|---|---|---|---|---|---|
LN | LA | VL | IL | Bio | RN | RL | RV | RSA | |
AT | *** | *** | *** | * | *** | *** | *** | *** | *** |
H | *** | *** | *** | *** | *** | *** | *** | *** | *** |
H x AT | * | *** | *** | ns | *** | ns | *** | *** | *** |
ns, *, **, *** Non-significant and significant at p ≤ 0.05, p ≤ 0.01 or p ≤ 0.001, respectively.
During the first 20 days of plant growth, leaf addition per plant displayed a linear response to harvest timing at all application timings (
Leaf area per plant displayed a quadratic response to harvest timing within the first 20 DAT (
Application timing | Leaf area m2·plant−1 | Biomass | MSRCE | ||||
---|---|---|---|---|---|---|---|
Leaf | Stem | Fibrous | SR | Total | |||
_________ g·plant−1 _____________ | % | ||||||
Untreated check | 0.9A | 42.5A | 69.5A | 6.8B | 268.8A | 390.7A | 86.6A |
0 DAT | 0.63C | 33.9B | 62.74A | 12.3A | 176.1B | 288.0B | 68.5B |
5 DAT | 0.72BC | 37.0AB | 64.02A | 7.4B | 190.7B | 302.0B | 70.4B |
10 DAT | 0.78AB | 41.8A | 64.22A | 7.1B | 267.2A | 383.A | 86.6A |
Means within rows followed by different letters are significantly different based on Fisher’s least significant difference mean separation test (P < 0.05).
At 20 DAT, untreated control plants had four and five adventitious and storage roots, respectively (
Sweetpotato marketable storage root conversion efficiency (MSRCE), defined as the percentage of marketable storage roots to total numbers of roots produced, did not differ between 10 DAT and the untreated check (87%) (
Additionally, peanut (Arachis hypogaea L.) grade and yield were not affected, probably because peanut has an indeterminate growth habit, which allows for compensation from early season stress like herbicide injury if given good growing conditions and sufficient recovery time. Also, Cardina and Swann [
Fresh storage root weight per plant, declined by 78% and 15% at 20 DAT, and 28% and 25% at 80 DAT for plants treated at 0 and 5 DAT, respectively, when compared to the untreated check (
At 80 DAT, stem dry biomass per plant ranged from 63 to 69 g and did not differ among application timings (
Plant biomass partitioned to leaves, stems, fibrous roots, storage roots and total roots at 80 DAT are presented in
Micrographs of transverse sectioned storage roots 20 DAT are illustrated in
Application timing | Leaf | Stem | Fibrous roots | Storage roots | Total roots |
---|---|---|---|---|---|
–––––––––––––––––––– % biomass –––––––––––––––––––– | |||||
Untreated check | 11 A | 15 B | 2 B | 72 A | 74 A |
0 DAT | 12 A | 22 A | 4 A | 61 B | 66 B |
5 DAT | 12 A | 21 A | 2 B | 64 B | 67 B |
10 DAT | 11 A | 17 B | 2 B | 70 A | 72 A |
Means within columns followed by different letters are significantly different based on Fisher’s least significant difference mean separation test (P < 0.05).
There were no meaningful differences between plants treated at 10 DAT and those of the untreated check. However, sweetpotatoes receiving S-metolachlor 0 or 5 DAT had reduced Chl a and b, total Chl, and carotenoid concentrations as well as reduced leaf area, root surface area, root volume, total lateral root length, fresh storage root, and marketable root weight, and total plant dry biomass. Findings from this study suggest that S-metolachlor applications should be delayed until 10 DAT to limit the herbicide’s potential impacts on sweetpotato growth, development, and yield.
This work was in part funded by the National Institute for Food and Agriculture, NIFA 2016-34263-25763 and MIS 043040 and the Mississippi Sweet Potato Council. We would like to thank David Brand for technical assistance and graduate students from the Environmental Plant Physiology lab for their help in taking measurements and processing samples. We thank Dr. K.N. Reddy, USDA-ARS, Stoneville, MS for providing the rainfall simulator, David Brand and Trevor Garrett for their technical assistance. This article is a contribution from the Department of Plant and Soil Sciences, Mississippi State University, Mississippi Agricultural, and Forestry Experiment Station.
The authors declare no conflicts of interest regarding the publication of this paper.
Abukari, I.A., Shankle, M.W., Reddy, K.R., Meyers, S.L. and Gao, W. (2019) Impacts of S-Metolachlor Application Timing on Sweetpotato Growth and Development. American Journal of Plant Sciences, 10, 780-795. https://doi.org/10.4236/ajps.2019.105057