Rust Fungi in Eastern Red Cedar Trees

Elizabeth Little, UGA Extension Plant Pathologist

Each spring, as the leaves of Rosaceous plants such as apple, pear, and hawthorn are emerging, the Eastern Red Cedar (Juniperus virginiana) produces the bright orange fructifications of a group of rust fungi in the genus Gymnosporangium. Early spring rains induce the cedar rust galls to break hibernation and produce gelatinous orange protrusions that release basidiospores. However, these basidiospores do not re-infect the cedar tree but instead drift off to find the appropriate secondary Rosaceous host (like apple, pear, and hawthorn). On the leaves and/or fruit of the secondary host, the fungus will mate and produce a different type of brightly colored spores. These spores only infect the cedar tree and this infection results in the cedar galls that can take up to two years to mature.

Cedar apple rust
Image 1 – Cedar apple rust (top of image), hawthorn rust and quince rust

The most common and visible cedar rust is cedar-apple rust (G. juniperi-virginianae). Cedar-apple rust forms large round galls on the small branches of the cedar trees. (see image 1) The basidiospores produced by the galls infect the newly emerging leaves of apple and crabapple. On susceptible cultivars, the resulting leaf lesions can cause the leaves to fall during years of heavy infection. Fungicide sprays may help on susceptible trees. However, in the home and/or organic garden, the use of apple cultivars with resistance to both cedar apple rust and fireblight is highly recommended. Removal of cedar trees in the immediate area can help but may not be practical. Check the Georgia Homeowner’s Pest Management Guide for recommended treatments and cultivars.

Two other cedar rusts are common in Georgia, quince rust (G. clavipes) and hawthorn rust (G. globosum). The perennial galls of quince rust are formed under the bark on the cedar branches and are nearly invisible until the gelatinous protrusions emerge from cracks in the bark (see image 1). The spores produced by these galls infect the fruits of the apple, quince, mayhaw, and pear (see image 2). Hawthorn rust produces small inconspicuous galls on the cedar (see image 1). This rust infects the leaves and shoots of ornamental hawthorn.

Quince rust on pear
Image 2 – Quince rust on pear fruit

Why are these annuals dying? What can I do?

Info taken from the UGA publication Crop Rotation and Cultural Practices Help Reduce Diseases in Seasonal Color Beds by Bodie Pennisi, Department of Horticulture and Jean Woodward, Department of Plant Pathology

Color bed disease fig 2 JWW-BPSeasonal color plantings can often appear healthy and beautiful one day and absolutely awful several days later. Total collapse at planting is not uncommon. Even though the chief culprit may be a particular disease, there are usually multiple causes at the root of the problem, all leading to plant demise.

Landscapers can reduce disease pressure in color beds by proper care:

  • Color bed disease fig 2 B JWW-BP
    Figure 2. Mixed planting and improper crop rotation in two consecutive years. The petunias are exhibiting signs of disease in the first year (upper photo) as well as the second year (lower photo).

    Choose plants suited to the sun and light conditions at the site. See this list of annuals with preferred growing conditions.

  • Prepare the soil in the beds to ensure both adequate water retention and drainage by incorporating several inches of organic matter each year.
  • Install annuals at a proper depth to avoid stressing root systems.
  • Remove old materials and spent blooms as much as possible to reduce disease inoculum.
  • Use water sensors on irrigation systems to provide water when it is needed, not on a specific schedule.
  • Choose water delivery methods, such as spray emitters directed at the base of plants, soaker hoses and drip irrigation, that will keep foliage dry.

Avoid planting the same species and cultivar of flowering crop, or crops belonging to the same family, in the same bed for more than two years. For example, petunias should not be followed with petunias because certain pathogens specific to petunias tend to accumulate and persist in the soil from year to year (Figure 2). Additionally, if ornamental tobacco is planted after petunia, it still could lead to plants being attacked by the same pathogens because the crops are closely related. This is of particular concern if the petunias were experiencing problems in the first year, which means that the pathogen is already in the soil and in sufficient quantity to cause damage.

Use the following list to select plants from different families to plant in successive years:

Common annual flowering species and their corresponding botanical family.
Family NameCommon Name of Flowers
AmaranthaceaeCelosia (Spike Celosia, Cockscomb), Alternanthera (Joseph’s Coat)
ApiaceaeParsley
ApocynaceaeVinca (Periwinkle)
AraceaeCaladium, Alocasia, Colocasia (Elephant Ears)
AsteraceaeAster, Marigold, Gazania, Mums, Sunflower, Black-eyed Susan, Echinacea, Zinnia, Mexican Sunflower, Melampodium, Ageratum
BalsaminaceaeImpatiens
BegoniaceaeBegonia
BrassicaceaeAlyssum, Ornamental Cabbage, Kale
CaryophyllaceaeDianthus
CommelinaceaeSetcresea (Purple Heart)
ConvolvulaceaePotato Vine (Ipomoea), Morning Glory
CrassulaceaeSedum
GeraniaceaeGeranium
GoodeniaceaeScaevola (Fan Flower)
LamiaceaeColeus, Salvia, Mints
LythraceaeCuphea (Cigar Plant)
PlantaginaceaeSnapdragon
PoaceaeGrasses (e.g., Pennisetum)
PortulacaceaePortulaca (Purslane)
RubiaceaePentas (Shooting Stars)
ScrophulariaceaeAngelonia (Summer Snapdragon), Torenia (Monkey Flower)
SolanaceaePetunia, Ornamental Tobacco, Ornamental Pepper, Browalia
VerbenaceaeVerbena, Lantana
ViolaceaePansy, Viola

Find more information:

Crop Rotation and Cultural Practices Help Reduce Diseases in Seasonal Color Beds

Flowering Annuals for Georgia Gardens

Flowering Bulbs for Georgia Gardens 

Native Plants for Georgia Part III: Wildflowers

Landscape Plants for Georgia 

Success with Mixed Containers Using Perennial and Woody Plants

Identification and Control of Spring Dead Spot

Alfredo Martinez, Turfgrass Pathologist, J.B. Workman, Graduate Assistant, Crop and Soil Sciences Department and Clint Waltz, Turfgrass Specialist

This Alert is an excerpt from the publication Identification and Control of Spring Dead Spot 

Spring dead spot Martinez
Figure 1 Multiple circular patches of dead, bleached grass are evident in the spring.

Spring dead spot (SDS) is a persistent and destructive disease of bermudagrass in Georgia. The disease is particularly prevalent and damaging in north Georgia, especially in the Piedmont region. However, SDS can be observed throughout the state after harsh winters and in areas where bermudagrass has been exposed to freezing temperatures for extended periods of time. The disease has also been observed in zoysiagrass, although less frequently.

Figure 2 Sharp edges between dead and healthy grass are observed once turfgrass greens up in spring.
Figure 2 Sharp edges between dead and healthy grass are observed once turfgrass greens up in spring.

Symptoms

As turfgrass “greens up,” well-defined circular patches of dead, bleached-out grass are noticeable in affected areas (Figure 1). Non-infected bermudagrass resumes growth, accentuating the infected areas. Sharp edges between dead and healthy grass are observed once turfgrass greens up in spring. (Figure 2).

Roots, rhizomes and stolons are sparse and dark-colored (necrotic) (Figure 3). Leaves become bleached, gray and straw-colored. Recovery from the disease is slow. Because the turfgrass in affected patches is dead, the primary means of recovery occurs by spread of stolons into the patch. Because recovery is dependent on lateral infill of surrounding bermudagrass, symptoms can remain visible well into the

growing season. If not managed properly, these patches may reappear in the same location the following spring along with weed species that may invade the voids. Patches can get larger year after year.

Disease cycle

The fungi casuing the disease are active in the fall and spring when cool, moist conditions exist. They do not kill bermudagrass directly; instead, they make turfgrass more susceptible to cold and freezing injury by feeding on roots, rhizomes and stolons.

Spread of these fungi primarily occurs through movement of infected plants or infested soil by equipment, people, animals and running water.

Infection of the turfgrass begins when soil temperatures are less than 70 °F. Typically, in Georgia, infection of susceptible grasses begins in late September or early October and will continue as long as soil temperatures are above 50° F. Fungal growth and plant infection can resume at these temperatures in early spring, coinciding with bermudagrass transitioning from winter dormancy (also referred to as “green up”).

Disease Control

Complete control of SDS in a single growing season is uncommon. It typically takes two to four years of proper cultural management and fungicide applications before acceptable control can be achieved. This has led to SDS becoming one of the more difficult diseases for growers to manage on an annual basis.

Resistant cultivars

The primary cultivars grown and used in Georgia (e.g., Tifway, TifSport, Tifton 10, Celebration, etc.) have shown susceptibility to SDS. However, SDS tolerance has been enhanced through breeding. Most “tolerant” cultivars (e.g., Patriot) may still get the disease but not as severely. In general, cultivars with more cold tolerance have less SDS than non-cold tolerant bermudagrasses. On sites where SDS has been a chronic problem, conversion to a tolerant cultivar is an option for disease management.

Cultural practices

Cultural practices that improve the cold-hardiness of bermudagrass can be particularly effective for managing SDS.

Since high nitrogen levels can reduce the winter hardiness of bermudagrass, apply no more than ½ pound of nitrogen per 1,000 ft2 be after mid-September.

Potassium applications in the fall (September or October) that total 1 pound of K2O per 1,000 ft2 can be helpful in improving the winter hardiness of bermudagrass and thus reduce SDS severity. Potassium applications should be applied based on soil test results.

A neutral to slightly alkaline soil pH can increase SDS severity. Maintain soil pH at 5.8 to 6.2. Use acid-forming fertilizers on sites with near neutral to alkaline pH. Apply iron, manganese and other micronutrients based on soil test results.

Any soil condition that reduces bermudagrass root growth such as compaction, excessive thatch (> ½ in) and poor drainage can also increase the severity of SDS. Core aeration and other practices that reduce soil compaction and encourage the production of new roots can be helpful in managing this disease.

Chemical Control

Timing, selection and application of fungicides are important for preventative management of SDS. Research has shown that one application of fungicide in the fall when soil temperatures are between 60° and 80° F provides the best control of SDS. When disease pressure is high, growers may want to make two applications. If a second application is necessary, it should be made four to six weeks after the first application when soil temperatures remain between 60° and 80° F. For complete meteorological information, see GeorgiaWeather.net . For improved results, it is recommended that fungicides be applied at high spray volumes (> 5.0 gal / 1,000 ft2) and/or immediately watered-in.

There have been mixed results from turfgrass managers around the state regarding chemical control of SDS. Those who have seen good results say they spray preventative fungicides that target SDS each year and have been doing so for several years. Therefore, it is important to keep in mind that controlling SDS takes time and control usually cannot be obtained in a single season.

A complete list of fungicides, formulations and product updates for SDS can be found in the annual Georgia Pest Management Handbook  and the Turfgrass Pest Control Recommendations for Professionals . Some fungicide options are exclusively for golf course settings. Always check fungicide labels for specific instructions, restrictions, special rates, recommendations, follow-up applications and proper handling.

This article is an excerpt from a more complete publication which can be found here.

What causes these irregular or circular patches in lawns?

Control Take-All Root Rot this Fall!

Alfredo Martinez, UGA Plant Pathologist and Willie Chance, UGA Center for Urban Agriculture

Take-all root rot -Clarissa Balibalian, Mississippi State Univ, Bugwood.org
Take-all root rot -Clarissa Balibalian, Mississippi State Univ, Bugwood.org

This year did your lawns show round or irregular dead or dying patches? Did the grass yellow or wilt even though the soil is moist? If so, these lawns may be infected with Take All root rot. This fungal disease affects cen­tipede, St. Augustine and Bermuda lawns

The fungus causing Take-All rots the lawn’s roots and aboveground runners (stolons).

To identify the disease look for:

  • Black, rotted roots.
  • Yellowed or dying areas of turf.
  • Stolons that are brown or black at the nodes or have dead patches.
  • One of the best ways to identify Take All is to look for the black, thread-like hyphae growing on the undersides of the stolons. You will need a micro­scope or a good hand lens to see these. Many UGA Extension County Offices have resources agents use to diag­nose diseases like this. Find your local UGA Extension office here.

This fungus prefers cooler weather – infecting lawns in the fall, growing through the winter and slowing growth in late spring. Much damage from this disease is done in the fall and spring. By the time we see disease symptoms (often in early spring and summer), the harm is done. Damage can be mistaken for green up problems. Expect the disease to be less active as temperatures increase.

Since this disease destroys roots, lawns may be slow to recover. Affected lawns are more susceptible to other stresses, like her­bicides and drought. Turf may not show evidence of the disease on the leaves until turf is stressed. For instance, a lawn with an unnoticed case of Take-All may be damaged or killed by the stress of a normal herbicide application. This can reflect poorly on the pesticide applicator.

Fall is the best time to control this disease. The best control is to improve cultural prac­tices to prevent the disease and to increase the vigor of the grass so that it will recover quickly. To slow disease progress:

  • Make sure the soil pH is not too high (Disease is less active below a pH of 6.5).
  • Water deeply and infrequently. Do not allow the soil to remain wet.
  • Remove thatch if the layer is thicker than one-half inch.
  • Mow at the proper height for your turfgrass.
  • Use fertilizers containing equal amounts of nitrogen and potassium.
  • Do not apply high amounts of nitrogen fertilizers in the fall. Typically warm season turf fertilization is completed by September 15.

Apply fungicides in September and again in October for best disease prevention. In warmer months, a fungicide may help, especially if sodding or plugging turf into affected areas. However, fall applications are best at controlling this disease. Fall applications should prevent the need for spring applications and should reduce Take-All damage in affected lawns and improve spring turf vigor.

For pesticide recommendations see the UGA publications –  Pest Management Handbook or Turfgrass Pest Control Recommendations for Professionals

For more information on controlling Take All see Turfgrass Diseases in Georgia or Enfermedades de los céspedes en Georgia

Fall Management of Large Patch Disease in Turfgrass

Large patch disease - Alfredo Martinez, UGA
Large patch disease – Alfredo Martinez, UGA

Alfredo Martinez, Extension Plant Pathologist

Large patch disease of turfgrass is most common in the fall and in the spring as warm season grasses are entering or leaving dormancy. Large patch is caused by the fungus Rhizoctonia solani. It can affect zoysia grass, centipedegrass, St. Augustinegrass and occasionally bermudagrass.

Symptoms of this lawn disease include irregularly-shaped weak or dead patches that are from 2 feet to up to 10 feet in diameter. Inside the patch, you can easily see brown sunken areas. On the edge of the patch, a bright yellow to orange halo is frequently associated with recently affected leaves and crowns. The fungus attacks the leaf sheaths near the thatch layer of the turfgrass.

Large patch disease is favored by:

  • Thick thatch
  • Excess soil moisture and poor drainage
  • Too much shade which stresses turfgrass and increases moisture on turfgrass leaves and soil
  • Early spring and late fall fertilization.

If large patch was diagnosed earlier, fall is the time to control it with fungicides. Consult the Pest Management Handbook , Turfgrass Pest Control Recommendations for Professionals  or your local Extension Office for fungicide recommendations. Fall fungicide applications may make treating in the spring unnecessary. Always follow label instructions, recommendations, restrictions and proper handling when applying pesticides.

Cultural practices are very important in control. Without improving cultural practices, you may not achieve long term control.

  • Use low to moderate amounts of nitrogen, moderate amounts of phosphorous and moderate to high amounts of potash. Avoid applying nitrogen when the disease is active.
  • Avoid applying N fertilizer before May in Georgia. Early nitrogen applications (March-April) can encourage large patch.
  • Water timely and deeply (after midnight and before 10 AM). Avoid frequent light irrigation. Allow time during the day for the turf to dry before watering again.
  • Prune, thin or remove shrub and tree barriers that contribute to shade and poor air circulation. These can contribute to disease.
  • Reduce thatch if it is more than 1 inch thick.
  • Increase the height of cut.
  • Improve the soil drainage of the turf.
  • Apply lime if soil pH is acidic (i.e. less than 6.0 – except on centipede lawns). Soil pH of more than 6.5 can encourage take all infections.

See the current Georgia Pest Management Handbook for more information. Check fungicide labels for specific instructions, restrictions, special rates, recommendations and proper follow up and handling.

Additional resources:

Turfgrass Diseases in Georgia or Enfermedades de los céspedes en Georgia

Turfgrass Diseases: Quick Reference Guide or Enfermedades de Cespedes Guia de Referencia Rapida

UGA Pest Management Handbook

Turfgrass Pest Control Recommendations for Professionals

Fungicide Efficacy Chart Available Online

Information from Jean Williams-Woodward, UGA Extension Plant Pathologist

I’ve been asked on numerous occasions for an efficacy table for fungicides labeled for ornamental plants. Well, myself, Alan Windham (University of Tennessee), Kelly Ivors (Cal Poly) and Nicole Ward Gauthier (University of Kentucky) put one together that lists products and their relative effectiveness for managing 14 diseases as part of a Southern Region IPM project. Diseases include:

  • bacterial leaf spots/blights
  • black root rot (Thielaviopsis basicola)
  • cedar rusts (Gymnosporangium rusts)
  • Conifer Tip Blights
  • Downy mildew
  • Fire blight
  • Fungal stem cankers
  • Fungal leaf spots
  • Fusarium stem rot
  • Passalora (syn. Cercosporidium, Cercospora) needle blight on Leyland cypress and other needled evergreens
  • Phytophthora root rot
  • Pythium root rot
  • Powdery mildew
  • Rhizoctonia blight/root rot

The table is not all inclusive, but it’s a start that we hope to expand upon and update. You can find the table here

Editor’s note – You can save the file as a pdf file to your computer. If you print it, do so in landscape format. I find the file to be more easily read as a pdf file on the computer since you can enlarge the size of the page. This is a great resource!

What is killing branches on this Leyland Cypress?

Bot canker - Dark, rust-colored dieback symptoms of Botryosphaeria (Bot) canker. G. Moody
Dark, rust-colored dieback symptoms of Botryosphaeria (Bot) canker. G. Moody

This disease is Bot canker. Bright, rust-colored branches and yellowing or browning of shoots or branches are the first observed symptoms. Closer inspection reveals the presence of sunken, girdling cankers at the base of the dead shoot or branch. Sometimes, the main trunk shows cankers that might extend for a foot or more in length. These cankers rarely girdle the trunk, but they will kill branches that may be encompassed by the canker as it grows. Read more info in the following publication including disease management.

Diseases of Leyland Cypress in the Landscape

See Entire Publication

Authors – Alfredo Martinez, UGA Plant Pathologist, Jean Williams-Woodward, UGA Plant Pathologist and Mila Pearce, Former UGA IPM Homeowner Specialist

Leyland cypress has become one of the most widely used plants in commercial and residential landscapes across Georgia as a formal hedge, screen, buffer strip, or wind barrier. The tree is best suited for fertile, well-drained soils. However, when young, the tree will grow up to 3-4 feet per year, even in poor soils. The tree will ultimately attain a majestic height of up to 40 feet.

Leyland cypress is considered relatively pest-free. However, because of its relatively shallow root system, and because they are often planted too close together and in poorly drained soils, Leyland cypress is prone to root rot and several damaging canker diseases, especially during periods of prolonged drought. Disease management is, therefore, a consideration for Leyland cypress.

This UGA Publication discusses several Leyland Cypress diseases and their management.

What is this common problem of centipede lawns?

Centipedegrass Decline

Alfredo Martinez, Extension Plant Pathologist and Clint Waltz, Extension Turfgrass Specialist
Adapted from original manuscript prepared by Drs. E.A. Brown, Retired UGA Extension Plant Pathologist and G. Landry, Retired UGA Extension Agronomist

Failure to green-up in the spring or successful green-up followed by decline and death in late spring and summer is a problem that can be encountered in centipedegrass-growing areas. Centipedegrass is subject to a condition called “centipedegrass decline.”

Many factors may contribute to this problem. It is important to be aware of these factors so that preventive and/or corrective steps can be taken. This problem can be prevented by proper management, which includes avoiding over-fertilization, preventing thatch accumulation, irrigating during drought stress (particularly in the fall), and maintaining a mowing height of 1 to 1.5 inches.

See the entire Centipedegrass Decline publication or read these sections:

Boxwood Blight Found in Georgia

Edited from an article by Jean L. Williams-Woodward, Extension Plant Pathologist. Read the entire article here.

Boxwood or Box Blight, caused by the fungus, Cylindrocladium pseudonaviculatum (syn. Cylindrocladium buxicola and Calonectria pseudonaviculata) has been confirmed in two residential landscapes in the Buckhead area of Atlanta.

The source of the introduction to one of the landscapes is unknown as new boxwood plants were not introduced into the landscape. The spores of the pathogen are very sticky and it is possible that the disease was introduced on worker’s tools or clothing. Plants within the second landscape were newly introduced from NC. Once introduced, the disease can be devastating to boxwood in landscapes and nurseries.

Boxwood Blight 2 JWW
Boxwood blight infected dwarf English boxwood in GA showing tan foliage and dieback. (Image by Jean Williams-Woodward)

Hosts: Dwarf English boxwood (Buxus sempervirens ‘Suffruticosa’) is highly susceptible and develops severe symptoms and rapid leaf drop. American or common boxwood (B. sempervirens) cultivars are also very susceptible. Cultivars of Littleleaf (Japanese) and Korean boxwood (B. microphylla and B. sinica, respectively) appear less susceptible because they don’t show severe symptoms and leaf drop, but they are still susceptible.

None of the commercial boxwood cultivars are immune to this disease. In fact, lesser susceptible (e.g. tolerant) cultivars may act as a ‘Trojan Horse’ introducing the disease into landscapes containing more susceptible cultivars.

The value of lesser susceptible cultivars is in the establishment of new boxwood hedges. If planting a new area, use a more tolerant cultivar to lessen your disease pressure in subsequent years. The disease also affects other plants within the Buxaceae family, including Pachysandra terminalis (ground spurge) and Sarcococca sp. (sweet box).

Boxwood blight JWW
Boxwood blight symptoms clockwise from upper left: Tan to gray leaf lesions with a darker purplish border on an English boxwood; Circular, tan spots with a brown border on upper leaves; Tan blighted leaves and bare stems on an infected plant; blackening of stems and browning foliage; and black stem lesions on bare branch tips. (Images by Jean Williams-Woodward)

Management: The best control is exclusion. Do not introduce the disease on infected plants or tools. Inspect all new boxwood plants for symptoms of the disease. Be sure to check the lower leaf canopy and interior stems. Keep new plants isolated and separate from existing boxwoods. Do not apply fungicides to plants in isolation that would mask symptom development. Monitor plants for at least four weeks prior to introducing them into existing plantings.

If Boxwood blight is detected, the infected plants and all fallen leaf debris needs to be bagged on-site and removed from the area to be buried in a landfill to prevent its spread. Transport plants in closed bags. Leaf litter blowing from open trucks could spread the disease to plantings along the roadway. Fallen leaf debris should be vacuumed and bagged, burned on-site or buried. Debris should not be composted. The fungus also produces microsclerotia (small clump of fungal hyphae) within roots and leaf debris of infected plants that allows the fungus to survive for years. Removal of existing garden soil and replacing with new soil is an option, but there is no guarantee that this will completely remove the pathogen.

Boxwood blight cannot be controlled with curative fungicide applications. Fungicides are only effective when applied preventively. Fungicide efficacy trials have shown that fungicides containing chlorothalonil (Daconil, Spectro, Concert II) and fludioxonil (Medallion, Palladium) provided the best control when applied preventively. To a lesser extent, fungicides containing azoxystrobin (Heritage), pyraclostrobin (Pageant), trifloxystrobin (Compass), and thiophanate methyl (Cleary 3336, OHP 6672) provided fair to good preventative control. Most are not labeled for use on either boxwood, Cylindrocladium or both; however, this is changing, so check labels. Remember, spraying plants after the disease is present will NOT control this disease.

Recommendations for Landscapers:

  • Inspect boxwoods on all properties. Look for symptomatic plants. As weather patterns become conducive (wet, humid, warm), disease symptoms may become noticeable and spread rapidly.
  • Submit suspect samples to the UGA Plant Disease Clinic in Athens through county extension offices for disease identification.
  • Train employees and clients on how to identify boxwood blight. Educate them on how easily the disease spreads.
  • Only purchase plants from nurseries that have a Boxwood Blight Compliance Agreement through their State Department of Agriculture. Many plants are brokered, so ask where plants were grown. Keep new plants in isolation and monitor for symptoms prior to installation.
  • Never install or prune or work in boxwoods when plants are wet.
  • Always visit non-infected landscape sites first. Move healthy to suspect diseased areas; never the other way around.
  • Disinfect pruners and other tools frequently within and between different blocks of plants within the same landscape, and especially when moving to different landscapes.
  • The best product is Lysol Concentrate Disinfectant (containing 5.5% O-benzyl-p-chlorophenol). Mix 2.5 Tbsp per gallon of water. This can be made and kept in spray bottles. Tools need to be wet for at least 10 seconds and allowed to dry to be effective.
  • A 10% bleach solution (1:9 part Clorox or 1:14 part Clorox Concentrate) for at least a 10 second soak can also be used, but this will oxidize tools. Soak and then let dry.
  • When leaving a site suspected or known to have boxwood blight, all tools, shoes, and clothing must be disinfected.
  • Get in the habit of wearing clean disposable booties or washing off debris and dirt entirely from soles of shoes between landscapes.
  • Changing and laundering clothes between sites would be ideal; however, it’s impractical. Wearing disposable paper pants is an option.

Find more information here

 

An unusual lawn invader appears during wet weather

Slime mold 3 turf disease pubThis is a Slime Mold growing on turf.

This information is taken from the UGA publication, Turfgrass Diseases in Georgia: Identification and Control 

Slime molds are caused by the fungi Physarum spp. and Fuligo spp.

All turfgrasses are susceptible to slime molds

Slime mold turf disease pubSymptoms: Large numbers of pinhead-sized fruiting bodies may suddenly appear on grass blades and stems in circular to irregular patches 1-30 inches in diameter. Affected patches of grass do not normally die or turn yellow and signs of the fungi usually disappear within 1 to 2 weeks. These fungi normally reproduce in the same location each year. The fungi are not parasitic, but they may shade the individual grass leaves to the extent that leaves may be weakened by inefficient photosynthesis.

Conditions favoring Slime Molds: Slime molds are favored by cool temperatures and continuous high humidity. An abundance of thatch favors slime molds by providing food directly in the form of organic matter.

Management:

Remove slime mold by mowing.

Remove using a gardening tool or high pressure stream of water.

For more information on slime molds and other turf diseases, see Turfgrass Diseases in Georgia: Identification and Control 

 

Slime molds elsewhere:

Dog vomit slime mold, Sandra Jensen, Cornell University
Dog vomit slime mold, Sandra Jensen, Cornell University

Other slime molds form irregularly shaped ‘blobs’ that grow on mulch, turf or other areas with organic matter. Read more about these in the publication, The Truth about Slime Molds, Spanish Moss, Lichens and Mistletoe

 

Also find pictures of various slime molds here.