Tag Archives: natural enemy

Cereal Leaf Beetle Biocontrol Project Underway

This month’s post is about a project being led by Jaime Cummings, the Field Crops and Livestock IPM Coordinator at NYS IPM. The goal is to improve biological control of the cereal leaf beetle, a pest of small grains. Before we tell you about the biocontrol project, you’ll need some background information on this pest and the other management options available. You can use the following links to navigate to each section of this post:

Cereal leaf beetles and damage they cause

Scouting for cereal leaf beetle and deciding when to spray

Biocontrol of cereal leaf beetle

Our project: Improving biocontrol of cereal leaf beetle

Cereal leaf beetle damage on the flag leaf of a winter wheat plant. (Photo by J. Cummings, NYS IPM)

Cereal leaf beetles and the damage they cause

The cereal leaf beetle (CLB), Oulema melanopus, can be a significant pest of winter and spring small grains production in NY, especially in parts of western NY.  This invasive species was first detected in Michigan in 1962, and has since become established in many grain producing states in the US, despite quarantine and pesticide eradication efforts in the 1960’s and 1970’s.

Left: a black beetle with a red middle (thorax), sitting on the leaf of a small grain crop; Right: a yellowish larva sitting on the leaf of a small grains crop
Figure 1. Cereal leaf beetle adult (A) and larval (B) stages. (Photos by J. Cummings, NYS IPM)

You may be familiar with this pest either in the larval or beetle stage (Fig. 1).  CLB has one or two generations per growing season, and the adults overwinter in hedgerows, woods or field margins.  We usually start seeing the adults move into small grains fields in April or May to lay eggs which develop into the damaging larvae.  The larger the larvae get, the more damage they inflict on the crop.  After about two weeks of feeding, the larvae drop to the ground and pupate for about two weeks before the adults emerge again.

When looking for these pests, keep an eye out for the typical larval feeding damage that looks like strips of green tissue missing between leaf veins.  Severely damaged leaves may appear skeletonized, and intense feeding pressure in a field may result in a ‘frosted’ appearance of flag leaves (Fig. 2).

green heads of winter wheat surrounded by leaves that have tan stripes on them
Figure 2. Severe cereal leaf beetle larval feeding on winter wheat. (Photo by J. Cummings, NYS IPM)

Considering that the top two leaves of the wheat/barley/oat crop are what contributes most to grain yield, severe infestations of CLB can significantly impact yield and grain quality.  Even in small grain or mixed stand forage crops, this pest can have negative effects on the yield and quality of the forage because they can significantly reduce leaf area and photosynthetic capability of the crop.

Scouting for cereal leaf beetle and deciding when to spray

It’s important to scout for this pest, usually starting in early to mid-June when larvae are first appearing.  The economic threshold for insecticide application for CLB is when you count an average of three or more larvae per plant before the boot stage or one or more larvae per flag leaf after the boot stage.  Occurrence of this pest can be inconsistent within a field, therefore plan to scout weekly and walk a random pattern throughout each field stopping at 10 random locations to count larvae on 10 plants at each location.  Because insecticides labeled for CLB target the larval stages, in order for your pesticide applications to be most effective, make sure that at least 25% of CLB eggs have hatched and that larvae are present and actively feeding when you decided to spray.  And, if you’re seeing adults in late June or beyond, it’s probably too late to spray for the larvae.  (Always follow label recommendations and restrictions when applying pesticides)

Paying attention to CLB populations in your fields via scouting is an important part of an integrated management approach for minimizing losses to this pest.  A growing degree day (GDD) model for CLB developed in Michigan determined that adult CLB begin to emerge around 350-400 GDD (base 48) to begin egg laying.

Biocontrol of cereal leaf beetle

Unfortunately, there is no specific host plant resistance available for CLB, but there are natural predators of the larvae and eggs which can help to keep the pest population in check, and possibly below the economic threshold when well-established in an area.  Lady beetles are known to prey on CLB larvae and eggs, and there is at least one egg parasite though it is not widely distributed.

There is also a CLB larval parasitoid wasp, Tetrastichus julis, which was originally introduced from Europe as a biological control agent in Michigan in 1967 (Fig. 3).  Subsequent releases into other states, including NY in 1973, have led to a sporadic establishment of this biological control parasitoid throughout small grain production areas of the US.

Tiny black wasp perched on dark brown larva clinging to a leaf
Figure 3. Tetrastichus julis, a parasitic wasp on a cereal leaf beetle larva. (Photo courtesy of Washington State Department of Agriculture)

Our project: Improving biocontrol of cereal leaf beetle

Given that CLB damage can be widespread and undermanaged in many small grains fields in NYS, and under the advice of Dr. Elson Shields (Cornell University Field Crops Entomologist), the NYS IPM program decided to try to determine the parasitism levels of CLB larvae in various locations around the state and to try to increase populations of the parasitoid in the Aurora area of Cayuga County, where the CLB tends to be a perennial pest.  The multiyear project was initiated this year, with CLB larval collections from locations in six counties.  However, there were no CLB present to collect at two of the locations, so the data collected in 2019 includes only four locations (Table 1).

Table 1.  Cereal leaf beetle collection efforts for determining parasitism levels in 2019.

Location County Collection date Crop # CLB larvae collected
Seneca Falls Seneca 6-Jun winter wheat, rye, barley 96
Aurora/Musgrave Cayuga 12-Jun spring barley 92
Ithaca Tompkins 12-Jun winter wheat, rye, barley 45
Penn Yan Yates 13-Jun spring oats and peas 110
Oriskany Oneida 11-Jun winter wheat 0
Homer Cortland 10-Jun winter wheat 0

 

At each location, a target of approximately 100 CLB larvae of all different sizes/growth stages were collected by hand from wheat, barley or oat fields.  The larvae were temporarily reared in incubation chambers on host plant leaves until approximately half of the larvae were dissected to determine baseline parasitism levels for each location (Fig. 4).

Left: Petri dishes with white filter paper and torn up leaves of oats; Right: Brown and yellow larvae of the cereal leaf beetle (some are squished) on a moist white filter paper in a petri dish
Figure 4. Cereal leaf beetle rearing chambers (A) and dissection process (B). (Photo by J. Cummings, NYS IPM)

The eggs of the parasitoid are visible when the CLB larvae are cut open under a microscope (Fig. 5).

close-up image of squashed yellow larvae. Dark head capsules are still visible, and small oblong eggs of the parasitoid can be seen next to one squished larva. The picture has the following labels: Dissected CLB larvae, and T. julis parasitoid wasp eggs from inside CLB larva
Figure 5. Dissected CLB larvae, and one with T. julis parasitoid eggs. (Photo by J. Cummings, NYS IPM)

After baseline parasitism levels were determined for each collection location, the other half of the CLB larvae were then released at the Cornell Musgrave research farm near Aurora, NY (Fig. 6).  This process will be repeated over the next few years.

Left: Hand hold an open petri dish filled with oat leaves, cereal leaf beetle larvae, and white filter paper; Right: Small dark larvae on an oat leaf with feeding damage
Figure 6. Cereal leaf beetle larvae with known level of parasitism being released in Cayuga County (Photos by J. Thomas-Murphy, Cornell University)

The goals of this project are to determine the established levels of the T. julis parasitoid around the state since the initial release in 1973, and to try to determine if we can increase its population at the research farm through consecutive releases.  From this first year of data collection, we know that the parasitoid population is low at the research farm in Cayuga County (6%) and at two of the collection sites (7% and 10%, in Seneca and Yates Counties, respectively), but was at approximately 30% at the Ithaca (Tompkins County) collection site (Fig. 7).

Graph shows that in Seneca County and Cayuga County only 7% and 6% (respectively) of cereal leaf beetle larvae were parasitized, while in Tompkins County the parasitism rate was 30%, and in Yates County the parasitism rate was 10%
Figure 7. Percent T. julis parasitized cereal leaf beetle larvae collected from various locations.

We also know that although there has been a need to spray insecticides to manage CLB at the research farm in Cayuga County and near the other collection sites, there has been no need to spray for CLB at the Ithaca (Tompkins County) collection sites.  It’s likely that the T. julis parasitoid population at the Ithaca site keeps the CLB population below economic threshold levels.  We hope that by intentionally distributing this parasitoid into an area with known CLB problems, we can establish a robust parasitoid population that may result in a reduction of necessary insecticide sprays for this pest.

This post was written by Jaime Cummings, Ken Wise, and Amara Dunn, all of the New York State Integrated Pest Management Program.

You too can ID aphids…and manage them with biocontrol

Lady beetle on a squash leaf, with a small cluster of pale green aphids nearby.
Caption: Lady beetles will eat any aphid species, but other aphid natural enemies are much more selective. (Photo credit: Amara Dunn)

Practicing good integrated pest management in the greenhouse requires correct identification of the pest. Accurate pest ID is also critical to successful use of biocontrol. Aphids are a good example. Biocontrol of aphids works best when you match the biocontrol agent to the aphid species you have. When I first learned this, I was a bit intimidated, because aphids are pretty small, and I’m not an entomologist. But the four aphid species you are most likely to encounter in your greenhouse are actually pretty easy to differentiate.

Anatomy of an aphid

In order to successfully ID aphids, you need to know (just a little) about aphid anatomy. All aphids are pretty small (between approximately 1/16 and 1/8 inches long). In addition to six legs and a body, aphids have antennae. Antennae attach near their eyes and are angled back over their bodies. They also have two little “spikes” that protrude from their rear end. These are called cornicles. Not so bad, right?

Diagram identifying the antennae on the aphid’s head and the cornicles attached at the rear of the abdomen.
Two features that will help you identify an aphid are the antennae attached to their head, and the two short cornicles attached to the rear end of their abdomen. (Diagram credit: Amara Dunn)

Green peach aphid

Enlarged photo of a light green, green peach aphid with identifying characteristics (indentation between antennae and cornicles that match the color of the body but with dark tips) labeled.
Green peach aphids vary in color from green to pink. Between their antennae you’ll see an indentation, and their cornicles are the same color as their body, with dark tips. (Photo credit: John Sanderson)

Green peach aphids come in different colors (from green to, well, peachy pink) and they are one of the smaller species. Their cornicles are the same color as their body (whatever that color is), and have dark tips on the ends. Green peach aphids also have an indentation in their head between the bases of their antennae.

Melon (or cotton) aphid

Enlarged photo of a dark green melon aphid with the distinguishing dark cornicles labeled.
Melon (also called cotton) aphids can be distinguished from green peach aphids by their dark cornicles. They also lack an indentation between their antennae. (Photo credit: John Sanderson)

Melon aphids (also called cotton aphids) also come in a range of colors that include light yellow, green, dark green, or almost black. Regardless of the body color, the cornicles will always be dark. Also, there’s no indentation in their head between the bases of the antennae. This is another small aphid species.

Foxglove aphid

Enlarged photo of a shiny green foxglove aphid with the distinguishing dark-spotted long antennae, dark leg joints, and darker green spots at the base of the cornicles labeled.
Foxglove aphids are shiny green with long antennae that look like they have dark spots on them. You’ll also see darker green spots at the base of each cornicle and dark leg joints. (Photo credit: Dan Gilrein)

Foxglove aphids are large (for an aphid). Their bodies are light green, but often shiny. There is an indentation in their head between their antennae. Their antennae are extra-long, extending well beyond the end of their body, and appear to have dark spots on them because the joints of the antennae are dark. The joints of their legs are also dark. Check where the cornicles attach to the body of the aphid. Foxglove aphids have darker green spots on their bodies at the base of the cornicles. These aphids usually like to hang out on the lower leaves of a plant, though they will infest flower petals sometimes.

Potato aphid

Enlarged photo of a potato aphid showing the segmented appearance and dark stripe running the length of the body.
Potato aphids have a dark stripe running down the length of their body, and they look faintly segmented. (Photo credit: John Sanderson)

Another large aphid, potato aphids come in pink and green. They look like they have a dark stripe running down the middle of their backs, and their body appears faintly segmented. They also have an indentation in their head between the antennae. Of the four species we’re discussing here, only the melon aphids lack this indentation.

To see these features, you will need a little magnification, but you don’t need a fancy microscope. Find a hand lens or a magnifier with 10X magnification. I like to keep one in my backpack so I’m always prepared.

Picture of a 10X hand lens. The lens folds out from a protective metal cover.
A 10X hand lens will enable you to magnify the features of an aphid that are important for identification.

There are even some relatively inexpensive 10X lenses you can snap on to your smartphone or tablet. Not only does this turn your device into a little microscope, but you can take a picture to document what you see (and show to an expert, later).

Picture of a macro lens that clips on to a smart phone or tablet.
Magnifying macro lenses can be clipped onto your smart phone or tablet, helping you both magnify and document the aphids you find.

You can also find (at least some of) these four aphid species outside. Last summer I spotted the aphid below on an acorn squash plant in August. Now that you know what to look for, what species do you think it might be?

Enlarged photo of a light green aphid with black cornicles. Do you know what species it is?
Use what you’ve learned to identify this aphid! (Photo credit: Amara Dunn)

One minor complication: Each of these four aphid species can either have wings, or be without wings. Usually aphids you find in a greenhouse have no wings, so you can stick with the above descriptions. But winged aphids can appear in the greenhouse, particularly when populations get very high. If you find aphids with wings in your greenhouse, the above descriptions won’t apply; ask for some help from your local extension office.

Enlarged photo of a winged (right) and non-winged (left) green peach aphid. The winged aphid is mostly black and looks very different from the pale green aphid without wings.
Both of the aphids in this picture are green peach aphids, but the one on the right has wings, and would be tricky to identify using the criteria described in this post. Get some help from an expert. (Photo credit: Whitney Cranshaw, Colorado State University, Bugwood.org)

Choosing the right natural enemy

Enlarged photo of a tiny parasitoid wasp surrounded by green aphids that appear to be covered in white wax. The wasp is inserting an egg into one of the aphids.
An Aphidius parasitoid wasp lays an egg inside its aphid prey. The developing wasp will kill the aphid. These happen to be cabbage aphids. (Photo credit: David Cappaert, Bugwood.org)

A good biocontrol option for aphids is a parasitoid wasp from the genus Aphidius. These tiny wasps are called parasitoids because they lay their eggs inside of aphids. As the young wasp grows, it kills the aphid and turns it into a mummy.

Enlarged photo of the tan shell of an aphid (mummy) with a hole in it, still sitting on a leaf
Aphids that are eaten from the inside out turn into dry, brown “mummies”. On this aphid mummy you can see the hole from which the adult wasp emerged. (Photo credit: Ken Wise, NYS IPM)

 

But if you want to purchase Aphidius wasps to release in your greenhouse (or the banker plants and prey that support them; read more here), you’ll need to know which Aphidius species to use. Aphidius colemani works well against green peach and melon aphids, while Aphidius ervi works well against foxglove and potato aphids. Another natural enemy you can use is Aphidoletes aphidimyza. This is a tiny fly whose larvae are voracious aphid predators. Although it seems to be less effective against foxglove aphid, it may work well in combination with another natural enemy.

An enlarged picture of what looks like a segmented worm (the larva of Aphidoletes aphidomyza), surrounded by pale green aphids
The larvae of the tiny fly Aphidoletes aphidomyza crawls around on leaves searching for aphids to eat. (Photo credit: Sarah Jandricic)

Like all biocontrols, Aphidius wasps and Aphidoletes larvae need to be released while your aphid population is very small, before it gets out of hand. Aphid infestations can explode very quickly! Scout your crop regularly, and keep records so you know which aphid species you are likely to have. (Consider the Pocket IPM Greenhouse Scout app to help you with your scouting and pest management.) Then plan your biocontrol releases accordingly. Parasitoids and predators for aphids should be released preventatively on crops that are prone to aphids.

If you’ve inspected your aphids at 10X magnification, and still aren’t sure which species you have, contact your local extension office for help with ID. If you are planning to send a picture, make sure that it is clear and shows the features of the aphid that you now know are important (antennae, body, cornicles).

You can learn more about aphid biocontrol in this factsheet from John Sanderson (Department of Entomology, Cornell University) on managing aphids in a greenhouse. Identification of these four common aphid species and which biocontrols you can use against them are also summarized here. The natural enemies listed in the chart are meant to be a starting place. Maximizing the efficacy of your aphid biocontrol program takes some trial and error and willingness to fine-tune your program to the crop and environmental conditions you’re dealing with. Suppliers of aphid natural enemies also have great information about how to use these biocontrol agents most effectively.

Happy aphid hunting!

This post was written by Amara Dunn and Betsy Lamb (NYS IPM) and John Sanderson (Entomology, Cornell University).

Creating habitat for beneficial insects: Starting Year 2

A close-up picture of a lady beetle on a plant
Lady beetle from our beneficial insect habitat plots

Last year I introduced you to the research field at Cornell AgriTech in Geneva, NY where Dr. Betsy Lamb, Brian Eshenaur and I are studying and demonstrating Christmas tree IPM. One part of this project is using perennial wildflowers to attract natural enemies of pests as part of an IPM strategy. The wildflowers (and some perennial grass) species were chosen because of the food and shelter they provide to pollinators. These same resources should make them useful to natural enemies of pests, too.

A plot containing wildflowers (some yellow and purple ones in bloom), with woodchip mulch visible between plants.
Mulching transplants planted in Spring 2018 was the most expensive establishment method, but these plots were looking pretty good a year later, even before we weeded.

By the end of our first field season, we had started using six different methods to establish wildflowers as habitat for beneficial insects (plus a weedy mowed control treatment). We also collected data on how much time and money we spent on establishment and how successful our weed management was. You can read about results from Year 1 in my post from last November.

But beneficial insect habitat establishment is not a one-year project. The establishment methods we started to implement in 2018 are ongoing, including periodic mowing of direct seeded plots, and hand-weeding of transplanted plots. We’ll keep track of how much time and money we invest in these plots in 2019, too.

The same plot is shown in two pictures. The picture on the left has some bare ground visible and many patches of grass and broadleaf weeds. The picture on the right shows the plot after it was mowed.
Plots that were direct seeded in 2018 will be mowed this year to favor the perennial beneficial insect habitat plants over annual weeds. This plot was treated with alternating herbicide and tillage during Summer 2018, and wildflower seed was planted in Fall 2018; (A, left) plot before mowing, (B, right) same plot after mowing.
Two pictures of the same plot before (left) and after (right) weeding. The un-weeded plot has lots of dandelion seed heads and no bare ground is visible. After weeding, you can see some bare ground and it’s easier to see the wildflower plants.
Plots that were transplanted in 2018 will be hand weeded this year to help the perennial wildflowers and grasses out-compete weeds. This plot was transplanted in Spring 2018 into bare (not tilled) ground and no mulch was used; (A, left) plot before weeding, (B, right) same plot after weeding.

And we want to know whether these plots are actually attracting beneficial or pest insects. So, in 2019 we are starting “Phase II” of our beneficial insect habitat work. We want to know which and how many insects (and other arthropods, like spiders) are being attracted to each type of plot. We will also count insects in no habitat plots (weedy, mowed occasionally) and mowed grass plots in the middle of the Christmas tree field for comparison.

Insect collection began in early May, and we are using four different techniques:

  • Sweep net – This is what it sounds like. We “sweep” a net through the air above the ground to capture mostly flying insects, or those who may be resting on the plants.
  • Butterfly and moth count – We walk through the field, counting how many of each butterfly or moth species we see in each plot.
  • Pan traps – These are bright yellow and blue bowls filled with soapy water. One bowl of each color is placed in each plot for 2 days, then we collect the insects that have been attracted to the colorful bowls and were trapped in the soapy water. This method will help us count flying insects, especially bees and wasps.

    A bright blue plastic bowl and a bright yellow plastic bowl are filled with soapy water and small rocks. Both are set on bare ground with some plants growing nearby.
    Bright blue and yellow bowls filled with soapy water and weighed down with rocks will attract certain flying insects. By counting insects collected in these pan traps, we can learn which insects are spending time in each plot.
  • Pitfall traps – These are clear plastic 16-oz deli cups (like you might use for take-out food) that are sunk into the ground in each plot. Insects that crawl along the ground fall in. We will use this method to count mostly ground-dwelling insects.

    A 16-ounce plastic deli cup sunk in bare soil of a plot so that the rim is level with the ground. The cup is half-full of liquid and also has caught a few green beetles. The trap is covered by a clear plastic dinner plate held about 6 inches above the ground by wire legs.
    A pitfall trap collects ground-dwelling insects. This one is protected by a rain cover. We didn’t want all the rain we’ve been getting this spring to overflow the traps and wash away the insects we caught!

I will write another blog post or two about this project during or at the end of this season. If you want to see more frequent updates, follow me on Twitter (@AmaraDunn). I’ll post weekly pictures of this project, including which beneficial insect habitat plants are blooming each week. You can also see lots of pictures from this project on Instagram (biocontrol.nysipm).

A new resource to help you protect pollinators

honey bee is perched on top of a young developing squash with the flower still attached
Many crops (and plenty of non-crop plants) rely on pollinators. Let’s protect them!

As I’ve discussed before, the natural enemies that provide biological control of pests include both larger creatures (like insects, mites, and nematodes) and microorganisms (fungi, bacteria, and viruses) that combat pests in a variety of ways. Microorganism natural enemies are regulated as pesticides (one type of biopesticide), while the larger natural enemies are not. Growers who are successfully using biocontrol insects, mites, and nematodes usually recognize that they need to apply pesticides in such a way that they are compatible with the biocontrol organisms they use. Take a look at my April post for a summary of online resources that can help you check compatibility of pesticides (including biopesticides) with natural enemies.

Some of these compatibility resources include information on the effects of pesticides (and biopesticides) on bees. Pollinators (including honey bees, lots of other bees, and some non-bees) are very important beneficial insects. You may have noticed that they have found their way into several of my blog posts. So, I wanted to let you know about a brand new resource (hot off the digital presses) to help you protect pollinators.

Image of the cover of the resouces entitled: Pesticide decision-making guide to protect pollinators in tree fruit orchards
“A Pesticide Decision-Making Guide to Protect Pollinators in Tree Fruit Orchards” is a terrific resource to help you choose pesticides (and pesticide combinations) that are least-toxic to bees.

A Pesticide Decision-Making Guide to Protect Pollinators in Tree Fruit Orchards” was written by Maria van Dyke, Emma Mullen, Dan Wixted, and Scott McArt. Although it’s focus is tree fruit orchards (and therefore the pesticides used in them), it should be useful for growers of other crops who want to choose pesticides that are least toxic to bees. A few highlights:

  • It includes information not only on pesticides used alone, but (when available) on synergistic effects when multiple pesticide active ingredients are used together. When you combine some chemicals (either in the tank or in the environment) the mixture is more toxic than both chemicals alone.
  • Where available, it summarizes pesticide toxicity to other bees besides just honey bees (e.g., bumble bees and solitary bees). You can read more about why this is important in this recent article.
  • It describes what we know about sub-lethal (in other words, negative effects on the bees that are less serious than death) effects of pesticides on bees.
  • It includes about half a dozen biopesticide active ingredients.
bumble bee feeding on a purple flower
Pollination is being done by more than just honey bees! This bumble bee (plus many more bee species) are important pollinators in NY.

You might be asking: If a chemical on this table is toxic to bees, will it also be toxic to the insect and mite natural enemies I am releasing or conserving on my farm or in my garden? I wish I had a definitive answer to that. As you can see from the nearly three pages of Literature Cited at the end of this document, collecting these data is a time-consuming process. For now, stick with the compatibility resources that are already available, and ask the companies you buy from (pesticides or natural enemies) about compatibility.

In closing, a huge amount of work went into this resource to summarize so much useful and current (as of October 2018) information in an easy-to-read table. Bravo to the authors! The Pollinator Network @ Cornell has other helpful resources for growers on protecting pollinators. Winter is a great time to make plans for using IPM and protecting the pollinators and natural enemies that are so good for the crops we grow!

Creating habitat for beneficial insects: Project update at the end of the first year

Fair warning, this is going to be a longer post. But partly that’s because there are so many pictures. I will start with the overview, then go a bit deeper into the weeds (literally and figuratively). To help you navigate more quickly, here’s a sort of table of contents that will quickly take you to the information you may be most interested to read:

Overview

Details on weed control

Timing of fall planting

The steps we followed to establish beneficial insect habitat

mixed wildflowers in a field
Some of our beneficial insect habitat plots looked really beautiful this fall! Others are still works in progress.

Overview
Remember back in June when I told you about the different techniques we were comparing for establishing habitat for beneficial insects? Time for an update! Here’s a brief, two-page summary of the first year of this project. For all the juicy details (and lots of pictures), keep reading!

First, remember that when I say “beneficial insects”, I mean both pollinators and natural enemies of pests. (Technically, arthropod would be a better term than insect, because spiders and predatory mites are some of the beneficial creatures we’d like to attract.) Fortunately, the same type of plants provide food and shelter for both pollinators and natural enemies on your farm or in your garden.

We used six different techniques to establish this habitat during Spring, Summer, and Fall of 2018. Treatment E was our control, where we did nothing but mow (after initial herbicide applications).

Treatment Fall 2017 Spring 2018 Summer 2018 Fall 2018
A Herbicide Herbicide, transplant  Weed 2x Replace dead plants
B Herbicide Till, transplant, mulch Weed 2x Replace dead plants
C Herbicide Till, direct seed Mow 3x Mow 1x
D Herbicide Till, plant buckwheat Mow 1x, till, plant buckwheat Mow 1x, transplant
E – control Herbicide Herbicide Mow 3x Mow 1x
F Herbicide Till, lay plastic Continue solarization Remove plastic, direct seed
G Herbicide Herbicide/till Herbicide 2x, till 1x Till 1x, direct seed

We transplanted the following species in treatments A, B, and D:

Common name Scientific name Number of plants in each 5 x 23 ft plot
Anise hyssop Agastache foeniculum 2
Common milkweed Asclepias syriaca 3
Blue false indigo Baptisia australis 2
Lanced-leaved coreopsis Coreopsis lanceolata 3
Purple coneflower Echinacea purpurea 2
Boneset Eupatorium perfoliatum 3
Wild bergamot Monarda fistulosa 2
Catmint Nepeta faassinii 2
Tall white beard tongue Penstemon digitalis 3
Black-eyed Susan Rudbeckia fulgida va. Fulgida 1
Little bluestem (grass) Schizachyrium scoparium 11
Showy goldenrod Solidago speciosa 1
New England aster Symphyotrichum novae- angliae 3
Ohio spiderwort Tradescantia ohiensis 2
NY ironweed Vernonia noveboracensis 2
Golden alexanders Zizia aurea 3

We planted seeds in treatments C, F, and G. The seed mixture we used was the Showy Northeast Native Wildflower & Grass Mix from Ernst Seeds, which included a more diverse species mix. This mix changes a bit from year to year. If you’re interested, you can learn about the details of the specific mix we used here.

 Labor and costs

Not surprisingly, there were big differences in how much time and money we spent on different treatments this first year. The costs and hours below are for a total area of 460 ft2 (0.01 A) per treatment. Most of the cost differences are due to the huge difference in seed versus transplant expenses. We paid about $2 per plant and needed 180 plants for each treatment. In contrast, we spent about $12.50 on seed for each treatment. You can find itemized lists of cost and time inputs for each treatment here.

Treatment Supply costs Time (person hrs)
A – spring transplant $417.12 13.2
B – spring transplant & mulch $539.29 20.4
C – spring seed $17.75 4.4
D – buckwheat & fall seed $390.55 10.3
E – control $2.32 2.6
F – solarize & fall seed $148.02 10.2
G – herbicide/tillage & fall seed $22.04 6.3

But, there were also big differences in how quickly the plants established. By September, both treatments (A and B) that had been transplanted in the spring looked like well-established gardens, with large, blooming wildflowers.

Transplanted plots with more weeds (left) and fewer weeds (right). Plot on the right had been mulched.
Four and a half months after transplanting, the beneficial habitat plants in treatments A (left) and B (right) were mostly growing well. But there was a big difference in weed control, in spite of similar amounts of time spent weeding each treatment.

We were generally pleased by how well most of the spring transplants survived. Although all the transplants came in 50-cell flats, some were larger than others, and the larger transplants survived better. We were fortunate to be able to plant into nice moist ground, so except for a little water on the day of transplanting, we didn’t irrigate. Survival might not have been as good if we’d had different planting conditions.

In contrast, the much less expensive treatment C was not looking too impressive even by October. A few partridge peas and blackeyed Susans bloomed this year, but otherwise it didn’t look much different from the control plots. In mid-summer, it looked like we were growing more ragweed than wildflowers.

Mower cutting ragweed (left) and mowed weedy mix with a few yellow flowers blooming.
In late July, it looked like we were growing mostly ragweed in treatment C (left). But after mowing four times during the summer and fall, you could definitely see the blackeyed Susans establishing (right).

Two of the treatments (F and G) were planted with seeds this fall, and one treatment (D) was transplanted this fall. So it’s really too early to tell how successful those treatments were. Stay tuned for more updates!

Mixture of mowed weeds with small plants (left), plot of bare ground (middle and right)
Fall transplanted (D) or direct seeded treatments (F and G) did not look very impressive by October 19, 2018. I’m curious to see what they look like next spring, summer, and fall!

Details on weed control

What about weeds? The graph below shows the average percent of the surface area of each plot that was covered with weeds versus planted beneficial habitat species on September 19, 2018. (Thank you, Bryan Brown, NYS IPM Integrated Weed Management Specialist for doing a weed assessment for us!) While we spent about the same amount of time weeding treatments A and B (the time difference is due to the time spent mulching treatment B), we achieved much better weed control with the mulch than without it!

Bar graph showing how much of each plot area was covered by weeds and by beneficial plants for each treatment
Bryan Brown assessed the percent of each plot that was covered with either the beneficial habitat species we had planted (blue) or weeds (orange). Each bar represents the average of four plots for each treatment, and the error bars show the standard error.

In treatment B, we spread chipped shrub willow mulch about 3 inches deep around the transplants. If I were to do this again, I would spread it thicker. I was disappointed with how many weeds were growing through the mulch just a month after transplanting.

mulched plot with lots of weeds on the left and fewer weeds on the right
The chipped shrub willow mulch we used was not as effective at suppressing weeds as I had hoped. On the left is part of the plot that had not been weeded yet. On the right is the part that was weeded on July 6. You can also see from this picture that there was a lot of lambsquarters in this field, and that we hadn’t been able to seed grass between the plots, yet.

But weeding twice during the season pretty much took care of the weeds in treatment B. Treatment A was also weeded twice, but as you saw in the graph earlier, weed control by the end of the season was not as effective.

Unmulched plot weeded (on the left), and before weeding (on the right)
Treatment A (transplanted in the spring, with no additional weed control) before (right) and after (left) hand weeding on July 6.

I think we’ll have to wait until next year to really understand how weed control is working in treatment C. Remember, the strategy was to slowly deplete the annual weed seedbank by allowing weeds to germinate, but preventing them from producing more seed. This is not supposed to be a quick establishment method, and it wasn’t.

blooming yellow flower (partridge pea)
This is what partridge pea looks like. It was one of the few species from the native wildflower and grass mix seeded in the spring (treatment C) that bloomed during this first growing season.

Buckwheat as a weed-smothering cover crop

By the time Bryan did our weed assessment, it had been 3 weeks since we mowed the second planting of buckwheat. Ideally, we would have transplanted shortly after mowing the buckwheat. But, the second crop of buckwheat was starting to set seed by the end of August, and our transplants weren’t scheduled to arrive until the end of September. So we mowed the buckwheat early to prevent it from contributing its own seed to the weed seedbank. But this meant that a lot of weeds had time to germinate before we transplanted the habitat plants. The buckwheat certainly suppressed a lot of weeds during the growing season, and I hope that this will help reduce weeds next year.

very young buckwheat seedlings in upper left, larger buckwheat seedlings in upper right, mowing mature buckwheat plants in lower left, cut buckwheat stubble in lower right
The buckwheat established quickly and crowded out many weeds. We mowed the first crop in July and re-planted. We had to mow the second crop about 3 weeks before we transplanted habitat plants (not ideal).

Solarization

Overall, we were pleased with how the solarization worked. We laid down 6 mil clear plastic (leftover from a nearby high tunnel) in early June, and did a little weed control around the edges of the plastic just once during the summer to prevent more weed seed production and to prevent shading of the plots.

Two people shovelling soil on top of clear plastic laid on the ground on the left, and on the right, clear plastic cover a small area of soil, with all the edges of the plastic buried
We laid 6 mil clear plastic over some plots on June 5, 2018 to solarize the soil underneath (and kill weeds).

We also learned that solarization will not control purselane. In contrast, the purselane thrived only under our clear plastic, and nowhere else in the field. The plot that had the most purselane also had the most other (mostly grass) weeds. I think the purselane pushed the plastic away from the soil and reduced the temperature a bit, allowing other weeds to grow.

clear plastic covering a small area of soil, but with weeds (purselane) growing underneath it
In some solarized plots, purselane grew happily under the plastic. Purselane was not a common weed anywhere else in the field during the season.

Some other plots were virtually weed-free when we pulled the plastic up in October. (Did you see how large the error bar was for weeds in treatment F in the weed graph above? This means there was a lot of variability between plots in this treatment.) Our soil temperature probe happened to be in the plot with the most purselane, and we still achieved maximum soil temperatures of 120 °F (at a depth of about 3 inches), compared to 90 °F in a nearby control (treatment E) plot.

Plots of ground that had been covered with clear plastic. On the left, very few weeds grew underneath. On the right, more weeds (purselane and some grasses) grew.
Solarization results were pretty variable from plot to plot. In some plots, it worked great (left). In other plots more purselane germinated and it didn’t work as well (right). We cut all the weedy vegetation off at the ground before direct seeding the beneficial habitat plants.

Repeated herbicide and tillage

At the weed assessment in September, the plot that had been alternately treated with herbicide and tilled looked best in terms of weed control. Like treatment C and all the treatments planted (by seed or by transplant) in the fall, I think we’ll get a better idea next year of how effective this method was at suppressing weeds.

A small plot of mostly bare soil with a few small weeds growing through. The plot is surrounded by grass
A few weeds were present a week after the last time the herbicide/tillage treatment (G) was rototilled. We broadcast, raked, and pressed beneficial habitat seed into these plots.

Timing of fall planting

One thing we struggled with this fall was deciding when to plant the wildflower and grass seed mixture. One source recommended the seeds be planted sometime between October and December. We were cautioned that if we planted the seed too early, some species (especially blackeyed Susans) might germinate this fall, and the young seedlings would be killed by an early frost before they established. But we were also afraid of waiting too long and not being able to till the soil (treatment G, only) if it got too wet. And we wanted a nice smooth seedbed. In treatment F, we suspected that leaving the clear plastic on into November would protect the weeds from the cooler weather. But we worried that taking it off too early would only allow more weed seeds to blow onto the bare ground.

person dressed in warm clothes sprinkling seed on bare ground. There is a field and a pond in the background.
We direct seeded October 18, 2018, after the weather cooled down a bit, and before the ground got too wet.

Finally, we compromised and planted the seeds on October 18 and 19, after our first hard frost, and once it looked like the nighttime temperatures would be in the 40’s (or below) for the next 10 days. It was only a week after the last tillage in treatment G, and the soil was still relatively dry. Those who live in the Finger Lakes know that late October and early November were pretty wet this year, so I’m glad we planted when we did. If you are trying to time fall seeding, I would recommend that you keep an eye on the 10 day forecast to see when temperatures are starting to cool. But if you get a dry sunny day to plant and it’s reasonably cool, I wouldn’t delay.

So if I want to plant habitat for pollinators and natural enemies next year, what should I do?

First, think about the time, money, and equipment you have available, as well as the area you’d like to plant. There probably isn’t a single right way to establish this habitat, but there may be a best way for you.

You can find more details on the techniques we used (and some links to other resources) here.

This post was written by Amara Dunn, Brian Eshenaur, and Betsy Lamb.

This work is supported by the Crop Protection and Pest Management Extension Implementation Program [grant no. 2017-70006-27142/project accession no. 1014000] from the USDA National Institute of Food and Agriculture.

Could your lawn use some biocontrol? Scout first.

white grubs found in a lawn
These white grubs can damage your lawn…but only if enough of them are present. Don’t waste time and money treating for them if you don’t need to!

For much of this summer, many people in NY had “water” at the top of their lawn care list. White grubs are another concern for home lawns. But finding a grub in your lawn does not automatically mean that you need to treat. Before you think about doing anything to your lawn to kill grubs, you should know how many grubs you have, and which species they are. If you do need to treat, consider using biocontrol.

Good news! It’s the perfect time of year to scout for white grubs. Starting in mid to late August, and continuing into October, grubs that hatched from eggs laid during the summer will be just beneath the surface of your lawn, feeding on the roots of your grass plants. This is the time to look for grubs. You might also notice some damage to your lawn from white grub feeding during this window.

life cycle of a white grub in your lawn
Late August through October is the perfect time of year to look for grubs in your lawn. The grubs will be young and close to the surface.

 

map of lawn and places you plan to sample for grubs
X marks the spot (to sample for grubs)! Select parts of your lawn where you think there might be grubs, sample in these places, and take notes on what you find.

Take a look at this fact sheet for detailed instructions on sampling your lawn. All you need is a piece of paper, something to write with, and a shovel or trowel. Check 1 foot by 1 foot squares around the lawn. If you have a bulb planter with a diameter of approximately 4.25 inches, or a golf course cup cutter, this works, too. Make notes about where you’ve sampled and how many grubs you found in each spot. Save the grubs from each sampling location separately.

peeling back turf from a 1 foot by 1 foot section and finding a grub
Cut a 1 foot by 1 foot square at each sampling location, and look for grubs in the roots of the grass. (Photo credit: NYS IPM)

If you didn’t find any grubs, please don’t treat your lawn! You are wasting money, and applying unneeded pesticides (or biocontrol nematodes) is never a good idea. If you did find grubs, it’s important that you determine which species they are. Why? Because the white grubs you are likely to find in NY are the immature (larval) stage of many different insect species. And each species causes different amounts of damage to your lawn. This means that the number of grubs your lawn can tolerate before it’s damaged – and therefore the number of grubs you should tolerate before treating for grubs – depends not only on the overall resilience of your turf, but also on the grub species you have. Check out the following table:

Number of grubs of each species before a treatment is justified
White grubs may look very similar, but they are not! Different species cause different amounts of damage to your lawn. If you find more than the number of grubs per square foot (or per 4.25-inch diameter soil core), you might consider treating your lawn for grubs. If not, you don’t need to do anything!

Fortunately, identifying grubs is easy, too! All you need is a penny, a hand lens with at least 15x magnification, and this online Grub ID tool. First, use the green “Learn how to identify grubs” button to find out which part of the grub to look at and how to hold it. Next, follow the instructions to compare each grub you found to the size of a penny.

online grub ID tool
The Grub ID tool explains exactly which part of the white grub to look at when you are identifying it. Just click the green button.
close-up picture of the rear end of a white grub, used in identification
Take a close look at the rear end of a white grub (using a 15x hand lens) to find out which species you have.

Finally, inspect its rear end with a hand lens to determine which species you have. Once you’ve identified the species, click on the species name to find specific management information. Now, look at that table again. For the species you found in your yard, do you have more than the listed number per square foot (or per 4.25-inch diameter soil core)? If not, then don’t waste time or money treating your lawn.

Most likely, only a few spots in your lawn (if any) warrant grub treatment. This is why you took careful notes about where you found grubs. Late August through September is also a good time to use a curative treatment for grubs in NY. The grubs are small and easier to kill. Some chemicals are effective when used at this time (but not the ones that are taken up by the plant!). A preventative pesticide that is taken up by the plant and kills the next generation of grubs when they start feeding in the late summer and fall should be applied in May or June. Before using any pesticide, find out if it is allowed in NY and find the product label using the New York State Pesticide Administration Database (NYSPAD). If a product isn’t listed in this database, you may not use it in NYS (even if you can buy it online). You must follow all instructions on the label.

Screen shot of the NYSPAD database search page
Use the product registration section of the New York State’s Pesticide Administration Database (NYSPAD) to check if a white grub pesticide is allowed in NY. When you get to the website, follow the three simple steps shown here.
Dark reddish brown grub (unlike healthy white grubs) killed by nematodes
Unlike the healthy white grubs you saw earlier in this post, this grub has been killed by nematodes.

But why use a chemical when you could use a biological control? Entomopathogenic nematodes are tiny beneficial worms that don’t harm plants, but kill grubs. See how they do it by watching this short video.  Why wouldn’t you want these nematodes working for you? Beneficial nematodes are a curative white grub treatment, so they should be applied between mid to late August and October. But you still only need to apply them to spots where grub numbers exceeded the thresholds in this table. You can purchase nematodes from garden centers or online garden supply stores. Look for the nematode species Heterorhabditis bacteriophora and Steinernema feltiae.

Dry, powdered form of nematodes used to kill grubs in your lawn
Grub-killing nematodes are sold as what looks like a dry powder. Dissolve them in non-chlorinated water (if available) before applying them to your lawn. Follow all package instructions.

For both species, make sure to follow the instructions on the package for storing and applying them. Nematodes will be harmed by ultraviolet light, so apply them around dawn or dusk, and water them afterwards to wash them into the root zone of the grass (where the grubs are). Any type of sprayer (as long as it doesn’t contain a fine mesh) or even a watering can will work to apply nematodes. If you use a sprayer, keep the pressure below 30 pounds per square inch. When you’re mixing up the nematodes, if non-chlorinated water is available, use that. Chlorinated water is fine for watering them in after you apply them.

Regardless of what treatment you use, scout your lawn again next year to find out how well your IPM strategy worked, and if there are other areas you need to treat (or not).

This post was written by Amara Dunn (NYS IPM) and Kyle Wickings (Department of Entomology, Cornell University).

Creating habitat for beneficial insects – early summer 2018 project update

Betsy, Deb, and Brian transplanting native wildflowers and grasses that will provide habitat for beneficial insects
Dr. Betsy Lamb, Deb Marvin, and Brian Eshenaur (left to right) transplanting native wildflowers and grasses on the edge of a research Christmas tree planting at Cornell AgriTech in Geneva, NY. These plants will provide food and shelter for pollinators and natural enemies of pests.

As I mentioned in my January post, I am excited to be working with two NYS IPM colleagues (Dr. Betsy Lamb and Brian Eshenaur) to demonstrate the costs, labor, and effectiveness of different methods for establishing habitat plants for pollinators and other beneficial insects. Remember, habitat for pollinators is also habitat for insects and mites that are natural enemies of pests on your farm or in your garden. Thus, planting for pollinators enables you to practice conservation biocontrol. These demonstration plots are located around a new research planting of Christmas trees at Cornell AgriTech at the New York State Agricultural Experiment Station in Geneva, NY. What we learn from this project can help you choose the best way to establish your own beneficial habitat (on your farm, around your home, near your school, etc.)

We are comparing 6 different methods of establishing habitat for beneficial insects, plus a control (Treatment E). Treatment E plots were sprayed with herbicide last fall and this spring, and will be mowed once this year. A summary of the plan for the other treatments is below.

List of treatments in this study. Each treatment is a different method for establishing habitat for beneficial insects
Comparing different methods for establishing plants that provide habitat and food for beneficial insects (pollinators and natural enemies of pests). Treatment E is the control.
seeds for plants that will provide habitat for beneficial insects
Native wildflower and grass seeds (A) were mixed with boiled rice hulls (B) to make them easier to broadcast (C). Much of what you see on the soil surface is just the rice hulls, but there are a few seeds that will hopefully grow into habitat for beneficial insects.

Because of when spring tillage occurred, plots that were scheduled to be tilled in the spring did not need a second herbicide application. About a week after spring tillage, Treatment C plots were direct seeded. I hand-broadcast a mixture of native wildflower and grass seeds at a rate of half a pound per 1,000 square feet. This worked out to be 26 g of seed for each 5-foot by 23-foot plot. To make it easier to broadcast such a small amount of seed, I first mixed the seed for each plot with about 3 cups of boiled rice hulls. After raking the seed in gently with a garden rake, I stomped the seed into the ground to ensure good contact with the soil. In a larger plot, you might use equipment like a cultipacker or lawn roller to achieve the same result.

 

young buckwheat plants
Two weeks (and three-quarters of an inch of rain) after seeding, buckwheat is establishing. It will hopefully crowd out weeds that would otherwise grow in these plots over the summer.

I broadcast (again, by hand) buckwheat seeds in the Treatment D plots at a rate of 70 pounds per acre (84 g for each of these small plots), and raked them in on May 31st. If the buckwheat establishes well, it will smother weeds during the summer, and we can mow and transplant into these plots in the fall. We plan to mow this crop of buckwheat when it starts flowering and then reseed it, for a total of two buckwheat plantings this summer.

 

We transplanted by hand 15 species of wildflowers and 1 grass species into plots assigned to Treatments A and B on June 4th. Because we were able to transplant right after it rained, it wasn’t too difficult to plant into the untilled plots (Treatment A). Some of them still had some stubble from the cover crops and weeds that had been growing in this field last year, and were killed by fall and spring herbicide applications.

Young wildflower and grass plants transplanted into untilled soil.
Native wildflowers and grasses transplanted into untilled soil. Some dead weeds and cover crop still remain on the soil surface.

The day after we transplanted into Treatment B plots, we mulched the plants to a depth of about 3 inches to (hopefully) control weeds for the rest of the summer while the habitat plants get established. We used chips from shrub willow because they were available, but other types of mulch would work, too.

wildflower and grass plants surrounded by mulch for weed control
These wildflowers and grasses will have help out-competing weeds from 3 inches of willow chip mulch.

Finally, we laid clear high tunnel plastic over the plots receiving Treatment F. Ongoing research from the University of Maine suggests that soil solarization can be an effective form of weed control, even in the northeast. So we’re giving it a try! To maximize the efficacy of this technique, we laid the plastic when the soil had been tilled relatively recently, and was still very moist. To keep the plastic firmly in place for the whole summer, we rolled the edges and buried them 4-5 inches deep, then stomped the soil down around all the edges. In the fall, we will hand broadcast a mixture of native wildflower and grass seeds over these plots (same mix as Treatment C).

a trench being dug around the edge of a plot to bury the edge of a sheet of clear plastic
Deb Marvin and Brian Eshenaur (left to right) dig a trench to bury the edge of this sheet of clear plastic. The goal is weed control by soil solarization.

We’ll give weed seeds in the Treatment G plots a few more weeks to germinate and grow (depending on the rain). Then we’ll kill them with an herbicide, and till these plots again to induce more weed seeds to germinate. Then we will repeat the herbicide application, till again, and so on. This should reduce the weed seed bank in the soil over the course of the summer. After a final tillage in the fall, we will broadcast seed from the same wildflower and grass mix we used for Treatment C. Fall is the recommended time for direct seeding beneficial insect habitat in the northeast. This treatment will also have the advantage of a full season of weed control prior to planting (also recommended). The downside is that it will take longer to establish the beneficial insect habitat.

As we get these plots established, we’re keeping track of the time spent on each treatment and the costs of materials. In the late summer or fall, Dr. Bryan Brown will assess weeds in each treatment, and I will photo document how well our beneficial insect habitat plants have established in each plot. All of these data will help you choose the method that fits your timeline, budget, and equipment/labor availability. Stay tuned for more updates…including an invitation to a field day (not this year), so that you can come see the results of this project for yourself!

Don’t make a mountain out of an ant hill – why ants in your lawn may not be a problem

Ant hill build in an area with short sparse grass
Ant hill built at the entrance to the nest. Cornfield ants (Lasius neoniger) prefer sunny areas with short and sparse grass. Photo courtesy of Matt Frye, NYS IPM.

Now that the weather is getting warmer and you’re spending more time outside, you might notice ant hills in your lawn. Reaching for a can of something that will kill them should not be your first move! These ants may be cornfield ants (known among scientists as Lasius neoniger). They are yellowish brown to dark brown, and about 1/8 of an inch long (or slightly longer). You are most likely to notice the ant hills they produce at the entrance to their underground nest in a sunny lawn where the grass is short and sparse (since this is their preferred nesting area). While the ant hills could be problematic on parts of a golf course where the grass must be kept very short, they aren’t big enough to be a problem in your backyard (if you’re mowing your grass to the correct height, which should be about 3.5 inches).

In addition to being harmless to humans – they don’t sting or bite – these ants are actually good for your lawn! They eat the eggs of grass pests, including Japanese beetles. One study found that when these ants were present in turf grass, they reduced the numbers of white grubs and other grass pest larvae. Choosing not to apply pesticides to kill these ants will help you practice conservation biocontrol in your own backyard! In other words, by protecting the natural enemies of lawn pests, you will have fewer lawn pests (and less damage) to worry about.

ants entering nest
Ants entering and exiting their nest. Cornfield ants (Lasius neoniger) are about 1/8 inch long, and range from light to dark brown. Consult an expert for correct identification. Photo courtesy of Matt Frye, NYS IPM.

Although these cornfield ants should be a welcome addition to your lawn for the reasons I’ve just described, if the hills they create are bothering you, there are some simple IPM solutions. Water and fertilize your lawn appropriately and use one of the top two height settings on your mower when cutting your grass. These strategies will help you achieve a denser, taller lawn. This type of lawn is less desirable for building new ant nests, and will make remaining ant hills less noticeable. For more information on maintaining healthy lawns, see the Cornell Turfgrass program’s Lawn Care: The Easiest Steps to an Attractive Environmental Asset.

A few final (but important) notes. Cornfield ants in your yard are a good thing. Ants in your home are a different story entirely, and NYS IPM has information on how to avoid in-home ant problems. If you are uncertain about what type of ant you have, consult an expert for proper identification. Your local extension office is a great place to start. Or, you can submit a sample to the Cornell Insect Diagnostic Laboratory.

Learn more about IPM for your lawn at Landscapes, Parks, and Golf Courses and ThinkIPM blog posts.

Mix and Match: Compatibility of biocontrol with other pest management strategies

Delphastus eating whitefly
This small black Delphastus is helping to control whiteflies in a greenhouse. It’s important that other pest management strategies in this greenhouse are used in such a way that they do not harm the Delphastus.

If you were going to tank mix chemical pesticides, you would of course read the label to check for compatibility before mixing products. The same concept applies when using living organisms for pest control. Whether you are using parasitoid wasps, predatory mites, microorganisms, or nematodes, you need to know whether your biocontrols are compatible with each other and any other pest management products you plan to use. For example, a biocontrol fungus might be killed if you tank mix it with (or apply it just before) a chemical fungicide. Insecticides (whether or not they are biological) could be harmful to natural enemy insects and mites. Even some beneficial insects are not compatible with each other because they may eat each other instead of (or in addition to) the pest.

It’s a good idea to keep an updated list of the products and organisms you plan to use for pest management, and their compatibility with each other. For biopesticides (remember the difference between “biopesticide” and “biocontrol”?), start by reading the label (see label excerpt below). You must follow all instructions you find there. Many manufacturers also provide lists, tables, databases, or apps to help you find compatibility information (some links at the end of this post). This is especially useful for insect, mite, and nematode natural enemies, which are not pesticides and do not have pesticide labels. When possible, obtain compatibility information from the manufacturer or supplier you will be using. Different strains of the same microorganism or nematode may have different sensitivities to chemicals.

Compatibility information from Preferal label
This excerpt from the label of the bioinsecticide Preferal provides some information on its compatibility with other products. All instructions on a pesticide label must be followed.

Remember that NY pesticide labels (including biopesticide labels) can be found through the NYSPAD system.

Below are some links to resources from several manufacturers and suppliers of biocontrol products. No endorsement of specific companies or products mentioned in this post is intended. If you know of a link to additional information that is missing, please let me know so that I can include it!

Beneficial nematodes from BASF – This chart describes compatibility of beneficial nematodes sold by BASF with natural enemies and pesticides. Note that only the genus name of each “biological” active ingredient is listed, and that over time, the names of some predatory mites (and whether they belong to the genus Amblyseius or Neoseiulus) have changed.

BASF table describing compatibility
Screen shot of part of a table summarizing compatibility of BASF beneficial nematodes with other pest management tools (in this case, biocontrol insects and mites).

Biobest Side Effect Manual – This side effects manual is available either as an interactive website, or as an app. Choose pest management products by active ingredient or name of the commercial product (including the biocontrol microorganisms Beauveria bassiana and several types of Bacillus thuringiensis). The list of “beneficial organisms” to choose from includes bumble bees and nematodes, but not beneficial microorganisms (fungi, bacteria, and viruses). Select active ingredients/commercial products and beneficial organisms from both lists, then use the legend to interpret the compatibility information that is generated.

Biobest compatibility information
Screen shot from the web-based version of Biobest’s compatibility resource.

 

 

 

 

 

 

 

Compatibility of BioWorks products – Compatibility sheets are linked from each product page.

BioWorks compatibility information
A portion of one of the compatibility sheets provided by BioWorks for their products.

 

 

 

 

 

Koppert Side Effects Information – This information is available either as an interactive website, or as an app. Select beneficial organisms of interest (by either the Koppert product name or the Latin name). Select one or more “Agents” (pest management products) by either the trade name or the active ingredient. Click on Results, and use the Legend to interpret the output.

Koppert compatibility information
Screen shot from the web-based version of Koppert’s compatibility resource.

If you plant it, they will come: Attracting natural enemies of pests

coreopsis flower (Coreopsis lanceolata)
This coreopsis flower (Coreopsis lanceolata) is more than just pretty; it also provides pollen and nectar for natural enemies to eat (when they aren’t eating pests!).

At this time of year, glossy catalogs start arriving in my mailbox full of pictures of all the beautiful fruits, vegetables, and flowers that I could grow after the snow melts. What these pictures don’t usually show are the arthropod (insect, mite, and related species) pests that can’t wait to eat what I plant. There are many IPM strategies you can use to fight back against these pests, and you can learn more here.

One of these strategies (and seldom is a single strategy sufficient) is to think about what else is growing near the vegetables, fruits, and flowers you want to protect. There aren’t just pest arthropods in your garden. These pests have natural enemies, too. If you provide good habitat for the natural enemies (including food and shelter), you will attract more natural enemies, and they are likely to consume more pests, protecting your plants. This is one way to practice conservation biocontrol – protecting and supporting the biocontrol organisms (natural enemies) that are already present.

So, what makes good habitat for natural enemies? In general, plants that bloom throughout the growing season (early spring to late fall) provide pollen and nectar to the natural enemies that use these as alternate food sources (in addition to pests). These plants also provide good shelter, both for natural enemies and the arthropods (including some pests) they feed on. As these natural enemies reproduce in the habitat you have created for them, they will also venture beyond this habitat and into your fruit, vegetable, and other flower plants, where they will eat more pests.

butterfly on purple coneflower (Echinacea purpurea)
This butterfly is finding nectar at a purple coneflower (Echinacea purpurea). Pollen and nectar are also important food sources for some natural enemies.

What is good habitat for natural enemies is also (in general) good habitat for pollinators. You have probably already heard how important pollinator protection is. Those glossy catalogs (or wherever else you buy your seeds or plants) likely sell species and varieties labeled as being “good for pollinators”. Just make sure you include plenty of variety. Because most plants (especially perennials) bloom for a limited time, you will need multiple species to ensure season-long blooms. Also, the variation in height and structure of the plants will provide diverse habitat for all of the different natural enemies you want to attract.

And what about protecting a larger area of plants (like a 5-acre field of pumpkins on a farm)? Will creating habitat for natural enemies help with pest control? The answer is complicated. It probably depends on a lot of things. How big the field is, how much habitat there is and where it’s located, which pests are a problem, and other pest management strategies (especially use of chemical pesticides) will have an impact. Research has shown that in some scenarios, yes, providing habitat for natural enemies can reduce some pest populations in some crops (one example).

Later this spring, I and two of my NYS IPM colleagues (Dr. Betsy Lamb and Brian Eshenaur) will set up a field experiment that will answer this question (over the next several years) in a Christmas tree planting. We will also compare different strategies for creating this habitat (seeds versus plants, and different weed control methods). Stay tuned for updates!

In the meantime, for suggestions on what flower species make good pollinator (and natural enemy) habitat, you can start by checking out lists of plants that provide good habitat for pollinators (also this one), or searchable databases of pollinator habitat plants. Your local Cornell Cooperative Extension office is another great resource. The Xerces Society also a resource on Habitat Planning for Beneficial Insects.