Landscape Alerts – Page 20 – Center for Urban Agriculture

Methods to Maximize Efficacy of Turfgrass Fungicides

September 24, 2015September 27, 2013 by Willie Chance

Alfredo Martinez, UGA Plant Pathologist

Weather conditions have been conducive for turfgrass diseases. We have received numerous calls and emails about proper control strategies, especially on the appropriate selection of turfgrass fungicides and their efficacy. Some ways to maximize the efficacy of turfgrass fungicides include:

Read carefully and follow the label directions before applying fungicide.
Apply fungicides at the rate specified on the label.
Always follow instructions for re-entry to the area.
Fungicides are not equally effective on all diseases. Proper fungicide selection is very important for disease management.
The best control is achieved by applying fungicides preventively (before disease is present).
Use compatible tank mixes at recommended label rates.
Use proper sprayer, nozzles and pressure to deliver appropriate coverage of fungicides. Flat fan or swirl chamber (raindrop) nozzles are recommended for turfgrass fungicide applications.
Avoid turfgrass stress (drought or temperature) before or at the time of application. This could interfere with maximum fungicide uptake, activity and efficacy.
Fungicides should be sprayed when air temperatures are between 60°F and 85°F (15°C and 29°C) for best results.
Fungicides should stay on the turfgrass foliage for at least 6 hours for most effective control. Delay mowing and other cultural practices as much as possible to give the fungicide a chance to work (for proper mowing frequency follow the one-third rule).
Use enough water when applying fungicides for adequate coverage. Usually 2.0 gal water/1000 sq. ft. should give adequate coverage and deposition. Some fungicides have to be watered-in for proper placement to ensure adequate activity.
Do not apply fungicides when conditions are windy to avoid drift and poor coverage. Wind velocity tends to be the lowest early in the morning and late in the afternoon.
Be patient if an application appears to have produced no results. Some fungicide application results can be seen months later.
Use fungicides judiciously and sparingly.

Some notes on Fungicide Resistance

Fungi sometimes develop resistance to particular fungicides, especially when a product is used repeatedly without alternating with chemically unrelated fungicides. When fungicide resistance develops, there is no value in increasing rates, shortening intervals between sprays, or using other fungicides with similar modes of action.

Fungicide resistance has been confirmed in a few instances for each of the following turfgrass diseases and fungicide groups:

Dollar spot against benzimidazole fungicides (thiophanate methyl) and DMI fungicides (propiconazole)
Gray leaf spot against strobilurin (QoI) fungicides (e.g. azoxystrobin, etc.)
Anthracnose against benzimidazoles (thiophanate methyl) and strobilurins (QoI) (azoxystrobin, etc.)
Pythium blight against phenylamide fungicides (mefenoxam)

Benzimidazoles (e.g. thiophanate methyl) and phenyl amides (e.g., mefenoxam) have the highest risk of resistance.

Strobilurins have a moderately high risk of resistance

DMIs and the dicarboximides (e.g. iprodione) have a moderate risk

Nitriles (e.g. chlorothalonil), aromatic hydrocarbons (e.g. PCNB), and dithiocarbamates (e.g. mancozeb) have a low risk of resistance.

Several general strategies are recommended to minimize the risk of fungicide resistance.

First, don’t rely on fungicides alone for disease control.
Avoid using turfgrass varieties that are highly susceptible to common diseases and follow good disease management practices.
Also, limit the number of times at-risk fungicides are used during a growing season and alternate at-risk fungicides with fungicides in a different chemical group (those with a different FRAC numeric code).
When using an at-risk fungicide, tank-mixing it with another fungicide from another chemical group (different mode of action) can also reduce the risk of resistance.

These are general principles that can help to reduce, but not eliminate risk. A fungicide-resistant pathogen population can still develop when these principles are practiced. Refer to product labels before tank-mixing products to ensure compatibility and to avoid phytotoxicity.

For major chemical group description, see the Georgia Pest Management Handbook – turf disease control section.

eLearn Urban Forestry Online Training

September 18, 2015September 25, 2013 by Willie Chance

The Office of the Southern Regional Extension Forester, the USDA FS Region 8–Urban and Community Forestry Program along with the Southern Group of State Foresters have partnered to design, develop and implement a state-of-the-art online, distance-learning program geared specifically toward beginning urban foresters and those allied professionals working in and around urban and urbanizing landscapes

To access the modules for free, please visit www.elearn.sref.info

To access the modules for International Society of Arboriculture and Society of American Foresters credit, please visit www.cfegroup.org

To access the modules for volunteer credit or a certificate of completion, visit www.campus.extension.org and look for the eLearn Urban Forestry–Citizen Forester course.

For more specific information, please contact Sarah Ashton, Educational Program Coordinator, Southern Regional Extension Forestry at sashton@sref.info.

Fall is the time to plant and transplant trees and shrubs

November 30, 2015September 23, 2013 by Willie Chance

By Merritt Melancon and Frank M. Watson , UGA Cooperative Extension

Although most planting and transplanting occurs in the spring, fall is the best time of year to plant or transplant trees and shrubs.

“Trees planted in the fall have an opportunity to establish an extensive root system while the plant is dormant,” said Frank Watson, the University of Georgia Cooperative Extension coordinator in Wilkes County. “The soil temperature in most parts of the state is warm enough to support root growth during most or all of the winter season.”

First make sure the trees or shrubs are healthy enough to plant in a new environment. If you’re buying new trees or shrubs from a nursery, make sure the trunk is not damaged, said Matthew Chappell, a UGA Extension nursery production specialist.

“If you see any damage to the bark, do not purchase (that tree),” Chappell said. The same goes for trees that are already on your property. You don’t want to stress an already damaged tree by transplanting.

Chappell added that picking trees with straight trunks and symmetrical canopies will save you a lot of heartache in the future. They’ll be easier to prune into a desired shape and typically are more structurally sound.

Also avoid purchasing pot-bound trees. Check the container for circling roots which indicate that the tree or shrub will have a poor root system after it’s been planted.

If you’re working with a tree that’s already on your property, help the plant take a break from producing new branches and leaves before transplanting. The plant can then put most of its energy into adapting to its new environment, not into producing new growth above the soil. Avoid applying high nitrogen fertilizers to plants for about two months prior to moving. Another way to reduce new growth is to restrict the amount of water applied. However, severe water stress prior to transplanting can weaken the plant and decrease the survival rate, Watson said.

In addition to having their growth restricted, transplanted shrubs and trees need to have their roots pruned. Pruning a tree’s roots — trimming them back until they fit inside the soil ball — maximizes the quantity of feeder roots that are moved with the plant. Ideally, plants targeted for fall transplanting would have their roots pruned the spring before they’re replanted, but they can still be pruned 30 to 60 days before transplanting in the fall.

Whether you’re working with a newly purchased plant or one on your property, it’s important to pay extra attention to preparing the plant’s new home. Properly preparing the planting site will affect root growth, which determines the plant’s chances of survival and subsequent growth.

The planting hole should be two to three times the diameter of the soil ball. Place the plant at the same soil depth it was grown at. If planting several small plants close together, it may be more efficient and better for the plant to prepare an entire bed.

When physically planting your shrub or tree, try not to disturb the soil ball of the plant. This will ensure maximum contact between the roots and the soil, which will speed the plant’s creation of its new root system.

A broken or loosened soil ball may prevent the plant from absorbing all of the water it needs. Wetting the soil around the shrub or tree can keep the soil ball together as you transplant. You may want to use wire baskets or other equipment that is available for moving plants.

Don’t plant trees and shrubs so that water pools on the surface of the planting hole. But remember, the plant will need extra water for the first two years.

Wait several months, maybe until the following spring, to fertilize the newly transplanted tree. This allows the root system to establish itself before spurring new growth above ground.

(Merritt Melancon is a news editor with the University of Georgia College of Agricultural and Environmental Sciences. Frank Watson is the University of Georgia Cooperative Extension agent in Wilkes County, Ga.)

Fall Interseeding and Overseeding: Not One and the Same

November 30, 2015September 16, 2013 by Willie Chance

Photo by Wayne Hanna, Professor, UGA Crop & Soil Sciences

Clint Waltz, Ph.D., UGA Turfgrass Specialist

It’s nearing that time again, time to think about fall interseeding and overseeding. These two practices are similar but technically not the same. Interseeding is the practice of seeding the same species into itself for the purpose of increasing stand density and recovery of lost grass. For example, tall fescue is interseeded into tall fescue in the fall to improve the overall stand which may have declined through the summer stress period. Different cultivars may be used but the turfgrass species remains constant. Adding centipedegrass seed to a thin centipedegrass lawn in the spring is another example where like species is seeded into like species.

Prior to interseeding, particularly with tall fescue, core aeration is a common practice. The benefits for core aeration are numerous (e.g. soil air exchange, relieving compaction, improved water infiltration, etc.), including improving a planting or seed bed conditions.

Overseeding, however, is the practice of temporarily introducing a second turfgrass species – typically a cool-season grass – into a permanent species – typically a warm-season species – for the purpose of winter color or traffic tolerance. An example of overseeding would be incorporating a second, or temporary, species into a permanent species, as in overseeding a bermudagrass baseball field in the fall with perennial ryegrass to have a green field in the early spring. The second species can compete with the permanent species for light, water, space, and nutrients, so overseeding can become an additional stress that has to be managed. Of the warm-season turfgrasses, bermudagrass is best adapted and tolerant of overseeding. It is ill-advised to overseed solely stoloniferous grasses like centipedegrass and St. Augustinegrass.

Successful overseeding involves growing healthy grass prior to overseeding, proper seed and seeding rate selection, overseeding timing and preparation, post planting maintenance, and spring transition. It is particularly important to maintain proper soil fertility, to relieve soil compaction, and to prevent excessive thatch development.

Overseeding selection involves selecting grasses that have characteristics suited to the particular needs. Annual ryegrass has been replaced by perennial ryegrasses, because of improved turf quality, color, stress and pest tolerance, and manageability. The “intermediate” ryegrasses tend to perform as the name implies somewhere between annual and perennial ryegrass, unfortunately most are more like annual ryegrass, not half way between the two.

Overseeding rates generally range between 5 and 10 pounds per 1000 ft² in lawns. In higher traffic situations, like sports fields and golf courses, the seeding range is between 8 and 12 pounds per 1000 ft². Using high quality “Certified” (blue tag) seed that is free of weed species is important to maintaining quality turf. It is also important to use seed treated with fungicides such as Apron particularly for early fall overseeding since seedling blight diseases can be a problem.

The ten pound seeding rate generally provides a rapid stand for fall use, while the five pound rate provides a thinner stand and may not provide much coverage until spring. Seeding rate generally relates to desired appearance and intended traffic or use. Higher trafficked areas need higher seeding rates. However, higher seeding rates may lead to more difficult spring transition. Balancing seeing rate with need and desired appearance is a management decision that can affect bermudagrass the following fall, so seed appropriately.

Proper timing of overseeding should result in a gradual transition from the warm-season turf to cool-season turf. Some common indicators that tell us it is time to overseed include: soil temperatures at a four‑inch depth approaching 75° F, night temperatures falling into the 50’s, average midday temperature in the mid-70° F, or 2 to 4 weeks before the average annual first killing frost date.

The objective to insuring a successful overseeding is good soil to seed contact. Seedbed preparations generally consist of close mowing or scalping, with some light vertical mowing, and blowing, sweeping, or vacuuming the loose plant debris from the soil surface. Generally, the more the turf is opened, the better the establishment rate, but the more competitive the cool-season turf will be in the spring. Seed which germinate in thatch or above the soil surface are more likely to dry-out and die before becoming established.

After dragging the seed into the soil, begin lightly irrigating to maintain good surface moisture and get the seed to germinate. This generally means irrigating three to five times per day until the seedlings are well established, but the total amount of water applied during a day would seldom exceed 0.5 inches. This irrigation practice should be done without causing puddling on the soil surface, free water encourages disease. After germination, gradually reduce the frequency and increase the time of irrigation until a normal irrigation program can be established.

Begin mowing when seedling height is 30% higher than desired. Use a mower with sharp blades and mow when the grass is dry to reduce seedling injury. Because first-mowed grass is tender and succulent a reel-type mower tends to lay seedlings over and not cut them. Using a rotary-type mower is commonly used for the first mowing with the fear of “ripping” seedlings being unfounded or insignificant. Transitioning to a reel-type mower after the second or third mowing can provide a high quality appearance.

Wait to fertilize after seedling emergence (generally three weeks after seeding) since earlier fertilizing may encourage warm-season turf competition. One pound of N per 1000 ft² per month is adequate with less (e.g. 0.25 to 0.75 lb N / 1000 ft²) commonly used. Use a soil test report to guide phosphorus needs but it is typical to apply some phosphorus shortly after seeding to improve rooting.

2013 reports show ryegrass and tall fescue seed crops being harvested earlier and faster than normal. This may not indicate a bountiful year for seed production. In fact, the forecast is for an average yield with a reduced quality due to more contaminants (i.e. weeds) in the seed fields. Combine these factors with short carryover inventories and reduced acres in production, the result may be higher seed prices for 2013.

Fall is rapidly approaching and seeding, be it interseeding or overseeding, will again be part of turfgrass management programs. Proper preparation prior to seeding and sound management afterward can provide a turfgrass surface that performs well and is attractive throughout the winter.

Emerald Ash Borer found with a foothold in North Georgia

November 30, 2015September 15, 2013 by Willie Chance

EAB Debbie Miller, USDA Forest Service — Emerald Ash Borer, Debbie Miller, USDA Forest Service, Bugwood.org

Sandi Martin and Merritt Melancon, University of Georgia

For years foresters and invasive insect experts have been on the lookout for the arrival of an unwelcome guest in Georgia. Now that it’s here, they hope the public will help restrict its spread within Georgia.

The small, iridescent-green beetle has killed millions of ash trees across a wide swath of Canada and the upper Midwest since it was first detected in 2002. The emerald ash borer — Agrilus planipennis— has spread south and west from infested areas over the last decade.

In July, researchers found adult emerald ash borers in survey traps in DeKalb and Fulton counties. A follow-up ground survey found larvae in nearby ash trees, confirming an established emerald ash borer infestation.

EAB larvae David Cappaert, Michigan State University — Emerald Ash Borer larvae in tree, David Cappaert, Michigan State University, Bugwood.org

Since 2005 University of Georgia invasive species experts have conducted an extensive trapping program in Georgia to screen for the emerald ash borer.

Georgia’s five species of native ash trees usually grow along stream banks. While ash only makes up about 1 percent of Georgia forests, they play an integral role in preventing the erosion of stream banks and keeping silt out of natural waterways. Ash is also a popular landscape tree, with 2.9 million trees planted around Georgia homes, businesses, parks and greenways. The value of these city trees in Georgia is estimated to be around $725 million.

EAB exit hole Penn. Dept. of Conservation & Natural Resources - Forestry Archive — Emerald Ash borer creates a D-shaped exit hole, Image from Penn. Dept. of Conservation & Natural Resources – Forestry Archive, Bugwood.rog

Although the adult beetle is an active flyer, it is believed that the primary way the beetle spreads is by hitching a ride on infested ash firewood, logs and nursery stock. Emerald ash borer larvae kill ash trees by burrowing serpentine tunnels in the inner layers of bark, preventing the tree from transporting water and nutrients to and from the tree canopy.

The Georgia Invasive Species Task Force will launch a public outreach plan to try to curb the spread of this pest in the near future. This task force consists of the Georgia Department of Agriculture, the Georgia Forestry Commission, UGA, the Georgia Department of Natural Resources and the USDA Animal and Plant Health Inspection Service.

People can help slow the spread of this beetle through Georgia by not moving firewood and by helping others to understand how dangerous it can be to move firewood from one area to another.

“To prevent the spread of emerald ash borer, it is important not to move firewood in which the insect can hide,” said Kamal Gandhi, associate professor in the UGA Warnell School of Forestry and Natural Resources. “Buy local firewood, whether camping or for your home.”

To help reduce the spread of the emerald ash borer in Georgia, homeowners with ash trees should have a certified arborist check their trees for signs of emerald ash borer infestations.

Suspected infestations should be reported immediately so that foresters or arborists can understand how the infestation is spreading. This will aid in the development of effective methods to reduce its spread and impact.

“The faster (scientists) can track the spread of the insect, the faster they can work to stop it,” said Joe LaForest, integrated pest management and forest health coordinator at the UGA Center for Invasive Species and Ecosystem Health.

The public can report suspected infestations by:

Using the SEEDN (Southeastern Early Detection Network) website www.eddmaps.org/southeast.
Using the SEEDN Smartphone App (available for iPhone or Android devices) apps.bugwood.org/seedn.html .
Emailing one of the Experts listed below; or by contacting a representative of the Georgia Forestry Commission, Georgia Department of Agriculture, the Department of Natural Resources or their local UGA Cooperative Extension agent.

For more information about the emerald ash borer and how to protect ash trees, visit www.gainvasives.org/eab.

For more information about spotting signs of emerald ash borer infestations, watch this.

(Sandi Martin is the public relations coordinator with the University of Georgia Warnell School of Forestry and Natural Resources. Merritt Melancon is a news editor with the University of Georgia College of Agricultural and Environmental Sciences.)

Other information:

Emerald ash borer factsheet

Emerald Ash Borer 2013 Update

Frequently asked questions about EAB in Georgia

Destructive tree pest discovered in Georgia

New bee advisory on neonicotinoid pesticides

November 30, 2015September 13, 2013 by Willie Chance

See original article from the Southern Region IPM News here

In an ongoing effort to protect bees and other pollinators, the U.S. Environmental Protection Agency (EPA) has developed new pesticide labels that prohibit use of some neonicotinoid pesticide products where bees are present. (This announcement affects products containing the neonicotinoids imidacloprid, dinotefuran, clothianidin and thiamethoxam. – Editor’s note)

“Multiple factors play a role in bee colony declines, including pesticides. The Environmental Protection Agency is taking action to protect bees from pesticide exposure and these label changes will further our efforts,” said Jim Jones, assistant administrator for the Office of Chemical Safety and Pollution Prevention.

The new labels will have a bee advisory box and icon with information on routes of exposure and spray drift precautions. Today’s announcement affects products containing the neonicotinoids imidacloprid, dinotefuran, clothianidin and thiamethoxam. The EPA will work with pesticide manufacturers to change labels so that they will meet the Federal Insecticide, Fungicide, and Rodenticide Act (FIFRA) safety standard.

In May, the U.S. Department of Agriculture (USDA) and EPA released a comprehensive scientific report on honey bee health, showing scientific consensus that there are a complex set of stressors associated with honey bee declines, including loss of habitat, parasites and disease, genetics, poor nutrition and pesticide exposure.

The agency continues to work with beekeepers, growers, pesticide applicators, pesticide and seed companies, and federal and state agencies to reduce pesticide drift dust and advance best management practices. The EPA recently released new enforcement guidance to federal, state and tribal enforcement officials to enhance investigations of beekill incidents.

More on the EPA’s label changes and pollinator protection efforts: http://www.epa.gov/opp00001/ecosystem/pollinator/index.html

View the infographic on EPA’s new bee advisory box: http://www.epa.gov/pesticides/ecosystem/pollinator/bee-label-info-graphic.pdf

Largest Mushroom Species in the Western Hemisphere Found Growing in Georgia

November 30, 2015September 12, 2013 by Willie Chance

Jeff DeLong, Undergraduate Researcher, Department of Plant Pathology
Marin Talbot Brewer, Assistant Professor, Department of Plant Pathology

See the original article here

Macrocybe titans is the largest gilled mushroom of any species in the Western Hemisphere, with caps in excess of 100 cm (3 ft). These giant mushrooms often occur in clusters within the tropics and subtropics of the New World, with previous sightings reported in the Caribbean, Central America, Mexico, South America, and only in Florida within the USA. Confirmation of this species in Georgia now extends the previously known range of M. titans. To our knowledge, this is the first report in Georgia. The mushrooms were discovered in October 2012 in the yard of an abandoned house in Athens. The fungus was identified as M. titans based on morphological features, habitat where it was found and DNA sequence similarity with known specimens.

The large, cream to buff-colored mushrooms were found growing in three large, dense clusters of eight to 10 mushrooms each (Figure 1). Single caps were 20-30 cm (8-12 in) across and convex with incurving at the margin. Younger mushrooms (Figure 2) had more convex caps than those of older specimens that became flattened with age. The smooth, dry surfaces of the caps were cinnamon-buff in color at the crowns fading to cream toward the margins. The inner flesh of the caps was white. The spore print, which was obtained by placing a cap on half white and half dark-colored paper and covering it with a bowl overnight, was creamy white (spore prints are necessary for proper mushroom species identification).

The stipes (stems) of the mushrooms were cream to buff measuring 30-45 cm (12-18 in) in length (Figure 3). They were club-shaped, tough and fibrous with bent-back scales and striations that curved around the stipe like stripes on a candy cane. The dense clusters of mushrooms were formed by the attachment of individual stipes at the base. Several miniature mushrooms less than 2.5 cm (1 in) in height were found attached to the base of the clusters. The cream to dull-yellow gills of the mushrooms were wavy, crowded and attached to the stipes. The species of the mushroom-producing fungus was confirmed as M. titans by sequencing a region of its DNA.

The mushrooms of M. titans, like the fruiting bodies of any fungus, are the reproductive structures where the spores are produced. The majority of the fungus is actually growing as filamentous hyphae underground and out of sight in decomposing dead and decaying plant matter in the soil. The three nearby clusters that were observed likely came from a single individual growing in the soil beneath the tall grass. M. titans is saprotrophic, which means that it decomposes dead or decaying plant matter and does not cause disease or rot on living grasses or trees.

It is not clear why or how this fungus found its way into Georgia. The mushrooms were close to landscape ornamental grasses and it may have arrived with the container plants, as Florida is a large producer of ornamental landscape plants. Or, it may be expanding its geographic range into Georgia due to climate warming or some other factor. This species was previously known as Tricholoma titans, but in 1998 was renamed Macrocybe titans, which literally translates from Latin into “giant large head.” The mushrooms are reportedly edible; however, never consume a wild mushroom without identification confirmation by an expert. Contact your county Extension office should you happen to encounter the giant mushrooms of M. titans.

We thank Kirk Edwards for discovering these mushrooms and contacting us about their identification.

References:

Bessette, A.E., Roody, W.C., Bessette, A.R., and Dunaway, D.L. 2007. Mushrooms of the Southeastern United States. Syracuse University Press, Syracuse, NY.

Bigelow, H.E., and J.W. Kimbrough. 1980. Tricholoma titans, a new species from Florida. Mycotaxon 11: 425- 429.

Pegler, D.N., Lodge, D.J., Nakasone, K.K. 1998. The pantropic genus Macrocybe gen. nov. Mycologia 90: 494-504.

Wood decay and falling trees are of great concern

November 30, 2015September 8, 2013 by Willie Chance

Jean Williams-Woodward, UGA Extension Plant Pathologist

Damage from past years of drought has taken a toll on our trees. Drought stress, construction injury, soil compaction and root girdling injures tree roots and provides an entry point for wood decay fungi. Storm damage, improper pruning, and wounding of trunks and branches also leads to wood decay fungi entry and infection.

Inonotus decay on base of oak tree — Inonotus dryadeus basidiocarps (conks) at base of oak tree. Tree should be removed due to its location and the potential hazard it poses. J. Williams-Woodward

Wood decay fungi can be classified into two broad categories: white-rot and brown-rot fungi.

White-rotters are those fungi that rapidly breakdown lignin and eventually cellulose. The infected wood becomes soft, spongy and stringy. It is usually a root and butt rot by a white-rot fungus that causes hardwood trees to fail and fall (see images of Inonotus dryadeus).

Brown-rotters are those fungi that breakdown cellulose and eventually lignin. Wood crumbles and shrinks. These rots are often referred to as a cubical or dry rot.

Regardless of the type of rot and what fungus is infecting the tree, if you see conks or mushrooms growing on the tree trunk or root flare, then the tree is in advanced stages of infection and decay and there is a concern for possible tree failure. It takes years for infection and growth of the fungus in the wood to produce outward signs of conks and mushrooms.

At present, methods to accurately detect how much rot is present in the wood do not exist. There are methods to detect internal cavities in trees to determine the thickness of a trunk shell, but measurements of spongy wood is difficult. There is no cure for wood decay. The best management approach is preventing injury to trunks, branches, and roots.

Forest Pest Insects in North America: a Photographic Guide

November 30, 2015September 4, 2013 by Willie Chance

R. G. Van Driesche¹, J. LaForest², C. Bargeron², R. Reardon³ and M. Herlihy¹

¹University of Massachusetts, PSIS/Entomology; ²University of Georgia; ³USDA Forest Service, State and Private Forestry

Orangestriped oakworm, Lacy L. Hyche, Auburn University, Bugwood

The photos present in this publication are intended to help foresters, urban landscaping employees, or others working with trees recognize some of the common pest insects affecting trees in North America and understand their life cycles and how they damage trees.

Prepare Now for Annual Bluegrass (Poa annua) Emergence this Fall

November 30, 2015August 29, 2013 by Willie Chance

Patrick McCullough, Extension Weed Specialist, University of Georgia

Annual weeds establish from seed and complete their lifecycle in one year. Summer or warm-season annual weeds (like crabgrass) establish in spring, grow actively in summer, and die out in fall. Winter or cool-season annual weeds (like annual bluegrass) establish in fall, grow from fall to spring, and complete their lifecycle in warm temperatures in late spring.

Failure to control annual weeds in late summer may predispose turfgrasses to winter weed infestations. In many lawns, it is fairly common to see turf with significant summer crabgrass populations have problems with annual bluegrass in fall. Open areas left in turf where crabgrass was once actively growing may permit annual bluegrass invasion during periods of peak seed germination. Controlling crabgrass now or in late summer could significantly improve turf cover, growth, and competition with annual bluegrass. See Table 1 for postemergence herbicide selection for crabgrass control in turf.

Late Summer Crabgrass Control Can Improve Annual Bluegrass Control This Fall

Quinclorac is a popular postemergence herbicide selection for crabgrass control in bermudagrass, zoysiagrass, and many cool-season grasses.

Single applications of quinclorac have excellent activity on mature, multi-tiller crabgrass plants at the seedhead stage in late summer. Bermudagrass, creeping bentgrass, perennial ryegrass, tall fescue, and zoysiagrass can be safely seeded seven days after a quinclorac application. Quinclorac requires the addition of an adjuvant, such as crop oil or methylated seed oil, for best results in established turf.

Mesotrione (Tenacity) can be used for postemergence crabgrass control in centipedegrass, perennial ryegrass, St. Augustinegrass, and tall fescue.

Mesotrione should be applied with a nonionic surfactant and will require two applications at a three week interval for late summer crabgrass control. These turfgrasses can also be safely established following mesotrione applications for crabgrass control. Currently, Tenacity can be used in nonresidential turf but will have residential lawns added to the label in the near future.

Fenoxaprop (Acclaim Extra) is a postemergence grassy weed herbicide for use in tall fescue, perennial ryegrass, and zoysiagrass.

Fenoxaprop has excellent activity on multi-tiller crabgrass with one application but efficacy is often reduced when crabgrass has seedheads present. Tall fescue and perennial ryegrass may be safely reseeded immediately after fenoxaprop applications. Late summer seeding of zoysiagrass is not recommended but newly plugged or sodded zoysiagrass may be treated with fenoxaprop.

Late Summer Cultural Practices to Reduce Annual Bluegrass Competition

Promoting turfgrass recovery from summer stress is critical to reduce annual bluegrass competition in fall.

Review cultural practices and make modifications if needed for lawns with crabgrass problems during summer months.

Mowing height significantly influences turfgrass competition with crabgrass, annual bluegrass, and other problem weeds.

Height of cut for most lawns should be no less than two inches. Raising the mowing height of tall fescue, for example, to three inches may significantly reduce annual bluegrass establishment in fall and reduce the need for postemergence herbicides in spring. Check mowing height for your turf-type.

Mowing frequency also influences turfgrass growth and susceptibility to annual bluegrass infestations.

Turf managers should mow lawns at least once per week during periods of vigorous growth to prevent scalping. Scalping thins out turf and may enable annual bluegrass establishment in open areas. While returning clippings is recommended to recycle nutrients to the soil, removal of clippings may be useful when annual bluegrass is present and producing seed heads. Removing clippings at this time will reduce the spread of viable seed through the lawn.

Encouraging turf recovery from summer stress may include modifications to fertilization programs.

Turf managers should consider reducing nitrogen fertilization during peak annual bluegrass germination and during periods of vigorous growth (cool weather). High nitrogen at these times encourages annual bluegrass spread and survival into winter and spring. Fertilizing dormant turfgrasses when annual bluegrass is actively growing will make these weed infestations worse.

Fall aerification of cool-season grasses may also influence annual bluegrass infestations.

Open areas of bare soil in turf following an aerification may encourage annual bluegrass infestations during periods of peak seed germination. Time aerifications in early fall to allow turf to recover before annual bluegrass germinates.

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