Annual Bluegrass Weevils

Diagnosis and Decision Making for Sustainable Annual Bluegrass Weevil Management

September 2019

Overview The annual bluegrass weevil (ABW), Listronotus maculicollis , is native to the Northeastern United States. ABW can have two to three generations per year and causes significant damage to golf courses in Northeastern and some Southeastern states. ABW adults overwinter in leaf litter, tall grasses, and other areas with dense organic matter that provide a buffer from harsh winter conditions. On a typical golf course, overwintering habitats include roughs, grassy native areas, forest edges, and tree and shrub lines. In spring — as early as March in upstate New York —ABW adults emerge from their overwintering areas and move toward golf turf playing areas to begin reproduction. This is when the ABW management season begins. For many reasons, management of ABW presents incredible challenges. First, the small size (about 1/8 inch in length) and cryptic nature of ABW adults make monitoring difficult. Second, as the life stages progress and ABW transitions through the egg and larval stages, observing ABW becomes increasingly difficult because most stages occur within the turfgrass stem. Complicating things further, as larvae mature, they emerge from the turf crown into the surrounding soil and proceed to feed on the crown and roots of the plant. Following the last larval development stage, larvae pupate below ground and the next adult generation emerges. This generation of ABW is more widely distributed on the golf course, and thus more difficult to find and diagnose than the previous generation. Finally, ABW development is highly asynchronous, meaning that the life stages of different individuals overlap, resulting in the presence of more than one life stage at a single time. Collectively, these factors make ABW a difficult insect to monitor and manage in an economically and environmentally sustainable way. However, by using proper scouting methods along with a well-informed decision-making process, you can improve the effectiveness and efficiency of ABW management at your facility. Traditionally, ABW management has focused primarily on scouting for and treating adults. However, to enhance control and to manage insecticide resistance in ABW, managers are encouraged to broaden their monitoring and management efforts to include ABW larvae in addition to adults. This publication provides the information needed to establish a successful ABW monitoring and management program.

Overwintering Adults Spring offers the best chance for controlling ABW adults before egg laying. The primary goals for adult scouting are to determine: • Timing and location of ABW emerging from overwintering sites —when and where on each course. Recording these location helps to narrow scouting efforts later in the season and future seasons. • Pattern of ABW adult movement following emergence toward short-mown turf. • Timing of peak activity of ABW adults, meaning the point at which the majority of ABW adults are found at or within the playing surface (fairway, tee, green) edge. In the Northeast, scouting for adults should begin as close to March 1 as feasible, though some colder areas may not have adult activity until late March or early April. It is also beneficial to install a degree day monitor on site where possible to help make your scouting efforts as efficient as possible.* In Northeastern states, ABW emergence should begin soon following snow melt or soil thaw and often corresponds with the full bloom stage of forsythia. The primary areas to scout for ABW include ones with historical ABW presence/ damage and their overwintering habitat of accumulated leaf litter and clippings such as turf adjacent to native areas/forest edges and shrub lines. Once begun, adult scouting should continue weekly until peak activity is observed at the nearest playing surface, noted, and these areas monitored later for larvae. This stage generally coincides with the phenological stage of half green/ half gold on forsythia and occurs at roughly 110 –120 growing degree days (GDD) – base 50 beginning March 1. Scouting Procedures– Adults General Practice Early in the season, sample in turf directly adjacent to known or likely overwintering sites to pick up on the emergence of overwintered adults. As spring progresses and once adults are found, begin sampling progressively away from overwintering sites, toward the closest short-mown playing surfaces (fairways, tees, greens) to track the movement of adults to these areas (Figure 1).

*Where helpful, degree day indicators associated with distinct ABW development stages have been included in this document.

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Method (Figure 3) • Mix soap into water and stir.

• Flag 3’ x 3’ areas where flushes will be conducted. • Spread the water evenly across the sampling area to fully saturate soil. • Let sit for 15 to 30 minutes. • Carefully scan surface of turf for insect activity. • Repeating the flush 5 minutes after the first will improve effectiveness.

Soapy flushes Soap flushing is a productive way to assess insect activity in soil and thatch. A dilute solution of lemon- scented soap acts as an irritant, flushing insects to the surface where they can be identified and counted. The procedure is inexpensive, but can be time consuming. Compared with other methods, soap flushing is somewhat limited in the amount of ground that can be covered. Soapy flushes are most effective when done above 10°C and effort should be made to conduct them at a similar temperature each time to improve the consistency of results. Materials (Figure 2) • Lemon-scented dish soap (2 tablespoons) • Bucket • Water (2 gallons) • Flags for marking (3’ x 3’ area) Figure 1. Direction of movement of ABW out of overwintering areas in early spring towards fairways.

Figure 3. Soapy flush procedure.

Figure 2. Supplies needed for conducting a soapy flush for ABW adults.

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Materials (Figure 4) • PVC pipe (2” diameter x 3’ long) with slits (0.5” wide x 10” long) cut in the pipe running lengthwise the full length of pipe. Evenly spaced 10” slits prevent pipe opening from collapsing. • PVC end cap to close one end of trap • 32 oz. and 8 oz. deli containers for trap catch • Soapy water or propylene glycol for killing trap catch • Plastic board for trap lid. This can be painted green to blend in with turf if installing at playing surface edge.

Linear pitfall traps These traps are a passive method for monitoring insect activity or movement. Traps allow for activity monitoring over time rather than at single points in time, but can be labor intensive to install and remove. They also require maintenance (debris removal from trap surface and emptying of trap contents). If done correctly, they are a good way to detect the first emergence of ABW adults and to track the duration of the emergence period.

Figure 4. Top – PVC (2” diameter x 3’ length) showing routed 0.5” linear slits for insect collection and end cap (left). Bottom – Fully set linear pitfall trap showing PVC trap tube and collection vial (right). Trap shows tops painted green for installation near playing surfaces.

Method • Trap should be installed by trenching a shallow strip of soil adjacent to likely overwintering area and burying the PVC pipe. Install pipe so that the entrance slits face up and only the top portion of the pipe shows above ground. • Collection container at end of trap can be installed using a cup cutter. Vacuum sampling While most expensive up front, vacuum sampling is easy to conduct and can be used across large areas of turf. Only minor modifications need to be made to the vacuum for insect sampling. As with other methods,

temperature will impact effectiveness and effort should be made to conduct vacuum sampling at a similar air temperature above 10°C at each event. Materials • Any standard leaf blower/vacuum • Insect collection insert for vacuum tube • This can be made out of a variety of durable and lightweight materials commonly found at any hardware or grocery store. Figure 5 highlights a few examples • For more detailed instructions on making a vacuum basket insert for your blower-vac, see the NYS BMP website publications page.

Figure 5. Different designs for vacuum basket. A – Thin- walled PCV pipe combined with aluminum flashing, rivets and adhesive foam rubber. B – Aluminum flashing and rivets. C – Large plastic jar with center cut out of lid. In all designs, the basket is cut on an angle to match that of the vacuum tube, and steel window mesh is fitted to the end of the basket to trap specimens.

A

B

C

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Method • Vacuum passes should be 30’ to 50’ in length depending on the size of the area being sampled. • Make linear vacuum passes parallel to overwintering areas or fairway edge, depending on time of season/ location of sampling (Figure 6). • At high throttle, walk slowly backward with vacuum chute and collection basket held flush to the turf surface for the full length of the sampling area. • Drop to idle, remove the vacuum basket, and empty contents via tapping the basket onto a sorting tray (Figure 7). • Sort materials on the tray for ABW adults. • It is best to avoid very wet turf when using this method.

Figure 6. Vacuum pass being taken midway between rough and fairway edge in late May.

Figure 7. Emptying and sorting vacuum basket contents onto sorting tray. Inset shows an adult ABW moving through the collected leaf litter.

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Decision Making and Management– Adults When deciding whether to treat adult ABW with pesticides, use scouting data to try to limit applications to only those areas where ABW are detected. Also keep in mind that poor timing and overuse of pyrethroids can result in insecticide resistance in ABW. If the decision is made to use a pesticide, be sure to time it carefully with peak adult activity at the playing surface. One week following pesticide applications, conduct follow-up vacuum sampling to gauge the effectiveness of the treatment and to make informed decisions Shortly after adults reach peak activity at the playing surface edge, they begin laying eggs within the turfgrass stem. This occurs ~175 GDD (base 50) , generally corresponds with the full bloom stage of flowering dogwood and is an important timeframe for the monitoring and/or management of ABW larvae. Preventive management of larvae. In many cases where ABW has proven problematic in previous seasons, it is wise to follow an adulticide application with a preventive application of a systemic pesticide such as products containing the active ingredient chlorantraniliprole, or imidacloprid in locations such as Long Island, NY, where chlorantraniliprole and other anthranilic diamides are prohibited. This application should correspond with peak ABW egg laying to allow time for the systemic products to be taken up by the plant so they are active upon egg hatch. Unfortunately, there are no easy scouting indicators for monitoring eggs, but managers can rely on their records of areas where ABW adults were detected along with both degree day and phenological indicators as noted above. Curative management of larvae. Shortly after egg laying is also when larval monitoring begins, and in most Northeastern states, this typically occurs in May. The goal of larval monitoring is to identify the time frame when most larvae transition from third to fourth instar and exit the crown of the plant, entering the surrounding soil and becoming vulnerable to many insecticides. This generally coincides with peak bloom for Catawba rhododendron (Rhododendron catawbiense) and ~350 GDD (base 50) . Larval sampling to monitor for this stage should occur weekly in areas where high adult activity was previously detected. about the need for later larval scouting. 1st Generation Larvae

Scouting Procedures– Larvae Soil Collection. Using a turf plugger, collect soil cores just inside the fairway edge in areas where high adult activity was detected. Collect one soil core every 2-3 feet across the area under investigation (Figure 8A). Backfill soil plug holes, remove cores from plugger (Figure 8B), bag and label. Cores should be processed as soon as possible and not be allowed to dry out.

Hand Sorting ABW larvae can be detected by hand sorting turf plants and soil plugs in the field. This method can be very effective in identifying when larvae begin to emerge from the turfgrass crown and inhabit the surrounding soil. However, it can also be labor- and time-intensive. Salt flotation This method generates the same high-quality data as hand sorting but allows for sampling from a greater area. Additionally, much of the method happens passively, meaning that larvae can be extracted while working on other tasks. The method also permits extracting young larvae from inside the turf plant as well as mature larvae inhabiting soil. Figure 8. Turf plug collection for monitoring of ABW larvae. A – Pattern of plug collection along fairway edge. B – Removal of soil from turf plugger.

Materials • Table salt • Tap water (lukewarm) • 16 oz. deli cups or similar container • Turf plugger • Divot mix for filling plug holes Method

• Make a salt solution using a ratio of 4 cups of table salt to 1 gallon of lukewarmwater. The solution can be made the day before to ensure that the salt dissolves completely.

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• Place turf plugs individually, turf side up, into 16 oz. deli cups (Figure 9A) then fill each cup with salt solution just until plug is submerged (Figure 9B). Thatchy plugs often float, exposing turf and soil, and reducing the efficiency of the method. If this happens, place a heavy steel washer or small stone on top of the plug to keep it submerged. • Larvae often appear within the first 30 minutes. However, plugs should be left in salt water for at least 90 minutes to account for variability in soil type and larva location, both of which can affect the speed of extraction. • Monitor the surface of the solution for ABW larvae. Larvae can be collected using the tip of a knife blade or other small implement and should be moved to a separate dish or paper towel for counting and life- stage determination.

good hand lens (~15x) and a penny. Place larva on the back of a standard penny, and use your hand lens to compare the width of the head capsule to the distance between two pillars on the Lincoln Memorial (Figure 10). The head capsule width on 1st through 3rd instar larvae fit easily between two pillars. The head capsule of the 4th instar larva, the stage at which larvae emerge from the turf crown, will just barely fit inside the gap between two pillars, and that of the 5th instar larva will be slightly larger than the gap between pillars. The goal of this scaling activity is not to measure every larva collected, but to develop a search image for gauging larval stages. Continue conducting weekly salt flotations until the majority of larvae recovered from your salt floats are 3rd to 4th instar. Using the general threshold of 1 larva per sample on average, along with the assessment of life stages and GDD will provide the best chance of accurately timing curative larvicide applications where necessary. Appropriate active ingredients at this stage include cyantraniliprole, indoxacarb, spinosad, and trichlorfon.

Figure 9. Salt flotation procedure. A – Turf plug being placed into deli cup. B – Turf plug flooded with saturated salt solution.

Decision Making and Management– Larvae The density of larvae that turf can tolerate varies. Typically, action thresholds range from 30 to 80 larvae per square foot for generation one. However, stressed turf may experience damage at much lower densities. A general threshold would be to use an average of one larva per sample, however, more precise timing of larvicides can be achieved by assessing the different larval stages. Remember, the goal is to identify the time frame when larvae are emerging from the turf crown and into the soil where they are vulnerable to curative pest management efforts. Measuring overall body size and head capsule width is the most accurate method for determining larval stage. This can be done using a

Figure 10. Standard U.S. penny with 1st through 5th larval instars (left to right). Compare width of the head capsule (amber – brown sphere at top of larva) to the width between two pillars on the Lincoln Memorial.

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